Coxiella burnetii) en fauna silvestre...
344
1 Epidemiología y control de la fiebre Q (Coxiella burnetii) en fauna silvestre ibérica. David González Barrio Tesis doctoral
Transcript of Coxiella burnetii) en fauna silvestre...
1
Epidemiología y control de la fiebre Q
(Coxiella burnetii) en fauna silvestre
ibérica.
David González Barrio
Tesis doctoral
2
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
3
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
4
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
5
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”
6
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.
7
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
8
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
9
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.
10
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
11
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
12
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
13
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,
14
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
15
(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,
16
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
17
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,
18
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).
19
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
20
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,
21
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
22
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
23
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
24
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).
25
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
26
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
27
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
28
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
29
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
30
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
31
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).
32
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
33
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.
34
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
35
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
36
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.
37
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
38
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
39
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
40
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
41
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
42
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.
43
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
44
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
45
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
46
(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,
47
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
48
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
49
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.
50
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).
51
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
52
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
53
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
54
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
55
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
56
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
57
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
58
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
59
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;
60
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
61
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,
62
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.,
63
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).
64
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
Artiodactyla Bovidae
Alpine ibex Capra ibex http://maps.iucnredlist.org/map.html?id=42397
France 5/61 (8.2) CFT Baradel et al., 1988
Switzerland 0/10 (0.0) PCR Marreros et al., 2011
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
Alpine chamois Rupicapra rupicapra http://maps.iucnredlist.org/map.html?id=39255 France
8/135 (5.9) CFT Baradel et al., 1988
n.a. (0.0-50.0) CFT Jourdain et al., 2005
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
European bison Bison bonasus http://maps.iucnredlist.org/map.html?id=2814 Poland
0/122 (0.0) SERO Salwa et al., 2007
0/40 (0.0) ELISA Rypula et al., 2011
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
Mouflon Ovis orientalis http://maps.iucnredlist.org/map.html?id=15739
Cyprus 23/74 (31.1) PCR Psaroulaki et al., 2014
23/77 (29.8) PCR Ioannou et al., 2011
Czech Rep. 2/2 (100.0) MAT Hubalez et al., 1993
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
Nilgai Boselaphus tragocamelus
http://maps.iucnredlist.org/map.html?id=2893 Poland 0/1 (0.0) ELISA Rypula et al., 2011
65
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Artiodactyla
Bovidae Sitatunga Tragelaphus spekii http://maps.iucnredlist.org/map.html?id=22050 Poland 0/1 (0.0) ELISA Rypula et al., 2011
Yak Bos mutus http://maps.iucnredlist.org/map.html?id=2892 China 75/552 (13.6) ELISA Yin et al., 2015
Cervidae
Moose Alces americanus http://maps.iucnredlist.org/map.html?id=818 Canada n.a. (16.5) IFA Marrie et al., 1993
Black-tailed deer Odocoileus hemionus colombianus
http://maps.iucnredlist.org/map.html?id=42393 USA 5/143 (3.5) ELISA Chomel et al., 1994
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
Fallow deer Dama dama http://maps.iucnredlist.org/map.html?id=42188
Czech Rep. 2/4 (50.0) MAT Hubalek et al., 1993
Hungary 0/22 (0.0) PCR Kreizinger et al., 2015
Italy 3/43 (6.9) CFT Giovannini et al., 1988
Spain 0/13 (0.0) IFA Ruiz-Fons et al., 2008
Sika deer Cervus nippon http://maps.iucnredlist.org/map.html?id=41788 Japan 42/61 (68.8) ELISA Ejercito et al., 1993
0/5 (0.0) IFA Neagary et al., 1998
Wapiti Cervus canadensis http://maps.iucnredlist.org/map.html?id=41785 USA n.a (22.0) CFT Enright et al., 1971
n.a. SERO McQuiston et al., 2002
Red deer Cervus elaphus http://maps.iucnredlist.org/map.html?id=41785
Czech Rep. 6/24 (25.0) MAT Hubalek et al., 1993
Slovakia 1/3 (33.3) PCR Smetanova et al., 2006
Spain
0/28 (0.0) PCR Astobiza et al., 2011
1/2 (50.0) PCR Ruiz-Fons et al., 2008
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
66
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Artiodactyla
Cervidae
Red deer Cervus elaphus http://maps.iucnredlist.org/map.html?id=41785 France 1/54 (1.8) CFT Baradel et al., 1988
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 deer
O. h. hemionus http://maps.iucnredlist.org/map.html?id=42393 USA 6/60 (10.0) ELISA Chomel et al., 1994
Roe deer Capreolus capreolus http://maps.iucnredlist.org/map.html?id=42395
France
25/222 (11.2) ELISA Candela et al., 2014
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
Poland 0/20 (0.0) ELISA Rypula et al., 2011
Czech Rep. 2/33 (6.0) MAT Hubalek et al., 1993
Slovakia 0/2 (0.0) PCR Smetanova et al., 2006
Spain
0/6 (0.0) PCR Ruiz-Fons et al., 2008
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
White-tailed deer Odocoileus virginianus http://maps.iucnredlist.org/map.html?id=42393 Canada n.a. (1.5) IFA Marrie et al., 1993
USA 112/624 (17.9) IFA Kirchgessner et al., 2012
Water deer Hydropotes inermis http://maps.iucnredlist.org/map.html?id=10329 Korea 13/196 (6.6) PCR Shin et al., 2014
18/196 (9.2) ELISA Shin et al., 2014
Suidae Eurasian wild boar Sus scrofa http://maps.iucnredlist.org/map.html?id=41775
Czech Rep. 2/32 (6.2) MAT Hubalek et al., 1993
France 0/209 (0.0) CFT Baradel et al., 1988
Germany n.a. (2.2-8.1) SERO Hening et al., 2015
Germany 6/220 (2.7) SERO Hartung, 2001
67
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Artiodactyla Suidae
Eurasian wild boar
Sus scrofa http://maps.iucnredlist.org/map.html?id=41775
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
Spain
1/1 (100.0) PCR Jado et al., 2012
1/125 (0.8) ELISA González-Barrio et al., 2015d
4/112 (3.6) PCR González-Barrio et al., 2015d
4/93 (4.3) PCR Astobiza et al., 2011
Wild swine USA 67/135 (50.0) SERO Clark et al., 1983
n.a. CFT Sidwell et al., 1964
Carnivora Canidae
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
Coyote Canis latrans http://maps.iucnredlist.org/map.html?id=3745 USA
n.a. (63.0) MAT Willeberg et al., 1980
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
Japanese racoon dog Nyctereutes procyonoides viverrinus
http://maps.iucnredlist.org/map.html?id=14925 Japan 0/30 (0.0) IFA Neagary et al., 1998
0/37 (0.0) ELISA Ejercito et al., 1993
Red fox Vulpes vulpes http://maps.iucnredlist.org/map.html?id=23062
Czech Rep. 1/1 (100.0) MAT Rehacek et al., 1977
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
UK 32/120 (26.6) ELISA Meredith et al., 2014
USA
n.a. (55.0) CFT Enright et al., 1971
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
68
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Carnivora
Felidae
Feral cat Felis silvestris catus USA n.a. CFT Enright et al., 1971
India n.a. (0.0) CAT Yadav and Sethi, 1980
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
Herpestidae Egyptian mongoose Herpestes ichneumon http://maps.iucnredlist.org/map.html?id=41613 Portugal 5/45 (11.1) PCR Cumbassá et al., 2015
Indian grey mongoose Herpestes edwardsii http://maps.iucnredlist.org/map.html?id=41611 India n.a. (0.0) CAT Yadav and Sethi, 1980
Mephitidae Spotted skunk Spilogale gracilis http://maps.iucnredlist.org/map.html?id=136797 USA
n.a. CFT Enright et al., 1971
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
Mustelidae
American mink Neovison vison http://maps.iucnredlist.org/map.html?id=41661 Spain 0/3 (0.0) PCR Astobiza et al., 2011
Eurasian badger Meles meles http://maps.iucnredlist.org/map.html?id=29673 Portugal 0/1 (0.0) PCR Cumbassá et al., 2015
Spain 0/74 (0.0) PCR Astobiza et al., 2011
Pine marten Martes martes http://maps.iucnredlist.org/map.html?id=12848 Spain 0/12 (0.0) PCR Astobiza et al., 2011
Sea otter Enhydra lutris http://maps.iucnredlist.org/map.html?id=7750 USA
0/103 (0.0) PCR Duncan et al., 2015
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
Least weasel Mustela nivalis http://maps.iucnredlist.org/map.html?id=14021 Portugal 0/2 (0.0) PCR Cumbassá et al., 2015
Spain 0/5 (0.0) PCR Astobiza et al., 2011
Otariidae Northern fur seal Callorhinus ursinus http://maps.iucnredlist.org/map.html?id=3590 USA
0/40 (0.0) PCR Minor et al., 2013
0/400 (0.0) PCR Duncan et al., 2014
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
69
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Carnivora
Otariidae Steller sea lion Eumetopias jubatus http://maps.iucnredlist.org/map.html?id=8239 USA 1/2 (50.0) PCR Kersh et al., 2012
Phocidae Harbor seal Phoca vitulina richardsi http://maps.iucnredlist.org/map.html?id=17013 USA /
Canada
17/27 (63.0) PCR Kersh et al., 2012
73/215 (34.0) IFA Kersh et al., 2012
Procyonidae Northern raccon Procyon lotor http://maps.iucnredlist.org/map.html?id=41686
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
Spain 15/48 (31.5) PCR González-Barrio et al., unpublished
USA 5/11 (45.4) SERO Randhawa et al. 1977
n.a. SERO McQuiston et al., 2002
Ursidae
Brown bear Ursus arctos http://maps.iucnredlist.org/map.html?id=41688 Croatia 0/13 (0.0) CFT Madic et al., 1993
American black bear Ursus americanus http://maps.iucnredlist.org/map.html?id=41687 USA 3/37 (8.1) CFT Dunbar et al., 1998
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
Viverridae Common genet Genetta genetta http://maps.iucnredlist.org/map.html?id=41698
Portugal 0/3 (0.0) PCR Cumbassá et al., 2015
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
Chiroptera
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
Phyllostomidae Greater spear-nosed bat
Phyllostomus hastatus http://maps.iucnredlist.org/map.html?id=17218 Fr. Guiana 0/17 (0.0) ELISA Gardon et al., 2001
Pteropodidae Indian flying fox Pteropus giganteus http://maps.iucnredlist.org/map.html?id=18725 India 2/14 (14.3) CAT Yadav and Sethi, 1980
Didelphimorphia Didelphidae Common opossum Didelphis marsupialis http://maps.iucnredlist.org/map.html?id=40501 Fr. Guiana 1/4 (25.0) ELISA Gardon et al., 2001
Gray four-eyed opossum
Philander opossum http://maps.iucnredlist.org/map.html?id=40516 Fr. Guiana 4/36 (11.1) ELISA Gardon et al., 2001
Diprotodontia Macropodidae
Agile wallaby Macropus agilis http://maps.iucnredlist.org/map.html?id=40560 Australia 1/5 (20.0) ELISA Cooper et al., 2013
Common walloroo Macropus robustus http://maps.iucnredlist.org/map.html?id=40565 Australia 1/3 (33.3) ELISA Cooper et al., 2013
Eastern grey kangaroo
Macropus giganteus http://maps.iucnredlist.org/map.html?id=41513 Australia n.a. (12.0) CFT Pope et al., 1960
70
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Diprotodontia
Macropodidae
Red kangaroo Macropus rufus http://maps.iucnredlist.org/map.html?id=40567 Australia n.a. (33.0) CFT Pope et al., 1960
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
Westerm grey kangaroo
Macropus fuliginosus http://maps.iucnredlist.org/map.html?id=40563 Australia
115/343 (33.5) ELISA Banazis et al., 2010
245/1017 (24.1) ELISA Potter et al., 2011
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
Erinaceomorpha Erinaceidae Southern white-breasted hedgehog
Erinaceus concolor http://maps.iucnredlist.org/map.html?id=40605 Iran n.a. (0.0) SERO/PCR Mostafavi et al., 2012
Eulipotyphla Soricidae
Crowned shrew Sorex coronatus http://maps.iucnredlist.org/map.html?id=29663 Spain 0/14 (0.0) PCR Barandika et al., 2008
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
Lagomorpha Leporidae
Brush rabbit Sylvilagus bachmani http://maps.iucnredlist.org/map.html?id=41302 USA n.a. (53.0) CFT Enright et al., 1971
European hare Lepus europaeus http://maps.iucnredlist.org/map.html?id=41280
Czech Rep. 12/263 (4.5) MAT Rehacek et al., 1977
Czech Rep. 0/23 (0.0) MAT Hubalez et al., 1993
Cyprus 15/31 (48.4) PCR Psaroulaki et al., 2014
Spain 2/22 (9.1) PCR Astobiza et al., 2011
European rabbit Oryctolagus cuniculus http://maps.iucnredlist.org/map.html?id=41291 Spain
0/6 (0.0) PCR Astobiza et al., 2011
176/464 (37.9) ELISA González-Barrio et al., 2015b
6/136 (4.4) PCR González-Barrio et al., 2015b
White-tailed jackrabbit
Lepus townsendii http://maps.iucnredlist.org/map.html?id=41288 USA n.a. CFT Enright et al., 1971
Japanese hare Lepus brachyurus http://maps.iucnredlist.org/map.html?id=41275 Japan 5/8 (62.5) ELISA Ejercito et al., 1993
Snowshoe hare Lepus americanus http://maps.iucnredlist.org/map.html?id=41273 Canada n.a. (49.0) IFA Marrie et al., 1993
11/22 (50.0) ELISA Marrie et al., 1986
Peramelemorphia Peramelidae Northern brown bandicoot
Isoodon macrourus http://maps.iucnredlist.org/map.html?id=40552 Australia 6/35 (17.1) ELISA Cooper et al., 2013
71
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Pilosa Bradypodidae
Pale-throated three-toed sloth
Bradypus tridactylus http://maps.iucnredlist.org/map.html?id=3037 French Guiana
1/1 (100.0) PCR Davoust et al., 2014
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
Rodentia Cricetidae
Bank vole Myodes glareolus http://maps.iucnredlist.org/map.html?id=4973
Czech Rep.
1/95 (1.0) MAT Rehacek et al., 1977
2/65 (3.0) CFT Syrucek and Raska, 1956
n.a. No ISO Syrucek and Raska, 1956
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
Brush mouse Peromyscus boylii http://maps.iucnredlist.org/map.html?id=16652 USA
1/2 (50.0) CFT Enright et al., 1969
24/78 (30.0) CFT Enright et al., 1971
5/306 (1.6) MAT Rieman et al., 1979
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
Common vole Microtus arvalis http://maps.iucnredlist.org/map.html?id=13488
Czech Rep.
0/31 (0.0) MAT Rehacek et al., 1977
1/82 (1.2) CFT Syrucek and Raska, 1956
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
North American deermouse
Peromyscus maniculatus
http://maps.iucnredlist.org/map.html?id=16672
Canada n.a. (76.1) PCR Thompson et al., 2012
USA
0/364 (0.0) CFT Stoenner et al., 1960
5/306 (1.6) MAT Rieman et al., 1979
6/24 (25.0) CFT Enright et al., 1969
72
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Rodentia Cricetidae
North American deermouse
Peromyscus maniculatus
http://maps.iucnredlist.org/map.html?id=16672 USA 72/291 (24.7)
CFT Enright et al., 1971
No ISO Enright et al., 1971
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
Dusky-footed wood rat
Neotoma fuscipes http://maps.iucnredlist.org/map.html?id=14587 USA
1/21 (4.7) MAT Rieman et al., 1979
2/31 (6.4) CFT Enright et al., 1969
9/311 (2.9) CFT Enright et al., 1971
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
European water vole Arvicola amphibius http://maps.iucnredlist.org/map.html?id=2149 Czech Rep. 1/6 (16.6) CFT Syrucek and Raska, 1956
n.a. No ISO Syrucek and Raska, 1956
Field vole Microtus agrestis http://maps.iucnredlist.org/map.html?id=13426
Czech Rep. 0/11 (0.0) MAT Rehacek et al., 1977
0/3 (0.0) CFT Syrucek and Raska, 1956
Poland n.a. ISO Tylewska-Wierzbanowska et al., 1991
UK 59/309 (19.1) ELISA Meredith et al., 2014
Gray dwarf hamster Cricetulus migratorius http://maps.iucnredlist.org/map.html?id=5528
Czech Rep. n.a. ISO Proreshnaya et al., 1960
Former USSR
0/1 (0.0) CFT Yevdoshenko and Proreshnaya, 1961
n.a. ISO Yevdoshenko and Proreshnaya, 1961
Southern marsh harvest mouse
Reithrodontomys megalotis
http://maps.iucnredlist.org/map.html?id=19410 USA
0/8 (0.0) MAT Rieman et al., 1979
3/28 (10.7) CFT Enright et al., 1971
3/28 (10.7) CFT Stoenner et al., 1960
n.a. No ISO Enright et al., 1971
Chihuahua vole Microtus pennsylvanicus
http://maps.iucnredlist.org/map.html?id=13452 USA 0/12 (0.0) MAT Rieman et al., 1979
73
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Rodentia
Cricetidae
Chihuahua vole Microtus pennsylvanicus
http://maps.iucnredlist.org/map.html?id=13452 USA
1/5 (20.0) CFT Enright et al., 1969
20/100 (20.0) CFT Enright et al., 1971
n.a. No ISO Enright et al., 1971
Musk rat Ondatra zibethicus http://maps.iucnredlist.org/map.html?id=15324
Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956
Former USSR
0/6 (0.0) CFT Yevdoshenko and Proreshnaya, 1961
n.a. No ISO Yevdoshenko and Proreshnaya, 1961
USA 2/19 (10.5) MAT Rieman et al., 1979
Northern grasshopper mouse
Onychomys leucogaster http://maps.iucnredlist.org/map.html?id=15338 USA 1/5 (20.0) CFT Stoenner et al., 1960
Pinyon mouse Peromyscus truei http://maps.iucnredlist.org/map.html?id=16694 USA
12/128 (9.3) CFT Enright et al., 1971
n.a. No ISO Enright et al., 1971
0/11 (0.0) CFT Stoenner et al., 1960
Revillagigedo Island ded-backed vole
Myodes gapperi http://maps.iucnredlist.org/map.html?id=42617 Canada n.a. (18.0) PCR Thompson et al., 2012
Dipodidae
Small five-toed jerboa Allactaga elater http://maps.iucnredlist.org/map.html?id=853 Czech Rep. n.a. ISO Proreshnaya et al.,1961
Woodland jumping mouse
Napaeozapus insignis http://maps.iucnredlist.org/map.html?id=42612 Canada n.a. (83.3) PCR Thompson et al., 2012
Echimyidae Cuvier's/Cayenne spiny rat
Proechimys cuvieri/ cayenne
http://maps.iucnredlist.org/map.html?id=22712110 Fr. Guiana 4/26 (15.4) IFA Gardon et al., 2001
Erethizontidae North American porcupine
Erethizon dorsatum http://maps.iucnredlist.org/map.html?id=8004 USA 0/3 (0.0) CFT Stoenner et al., 1960
0/6 (0.0) MAT Rieman et al., 1979
Gliridae
Hazel dormouse Muscardinus avellanarius
http://maps.iucnredlist.org/map.html?id=13992 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956
Garden dormouse Eliomys quercinus http://maps.iucnredlist.org/map.html?id=7618 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956
Spain n.a. ISO Perez Gallardo et al., 1952
Heteromyidae Houserock chisel-toothed kangaroo rat
Dipodomys microps http://maps.iucnredlist.org/map.html?id=42603 USA 5/189 (2.6) CFT Stoenner et al., 1960
n.a. ISO Stoenner et al., 1960
74
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Rodentia
Heteromyidae
Owyhee river kangaroo mouse
Microdipodops megacephalus
http://maps.iucnredlist.org/map.html?id=42606 USA 0/2 (0.0) CFT Stoenner et al., 1960
Great-basin pocket mouse
Perognathus parvus http://maps.iucnredlist.org/map.html?id=42610 USA 0/14 (0.0) CFT Stoenner et al., 1960
0/22 (0.0) MAT Rieman et al., 1979
Heermani's kangaroo rat
Dipodomys heermanni http://maps.iucnredlist.org/map.html?id=42600 USA
0/17 (0.0) MAT Rieman et al., 1979
0/2 (0.0) CFT Enright et al., 1969
1/59 (1.7) CFT Enright et al., 1971
n.a. No ISO Enright et al., 1971
Long-tailed pocket mouse
Chaetodipus formosus http://maps.iucnredlist.org/map.html?id=4331 USA 0/47 (0.0) CFT Stoenner et al., 1960
Ord's kangaroo rat Dipodomys ordii http://maps.iucnredlist.org/map.html?id=6691 USA 16/312 (5.1) CFT Stoenner et al., 1960
n.a. ISO Stoenner et al., 1960
Muridae
Lesser bandicoot rat Bandicota bengalensis http://maps.iucnredlist.org/map.html?id=2540 India n.a. ISO Yadav and Sethi, 1980
Black rat Rattus rattus http://maps.iucnredlist.org/map.html?id=19360
Czech Rep. 1/15 (6.6) CFT Syrucek and Raska, 1956
n.a. No ISO Syrucek and Raska, 1956
Fr. Guiana 0/17 (0.0) ELISA Gardon et al., 2001
Netherl. 0/56 (0.0) ELISA Reusken et al., 2011
5/166 (3.0) PCR Reusken et al., 2011
Brown / black rat R. norvegicus/R. rattus Cyprus 32/136 (23.5) PCR Psaroulaki et al., 2014
63/494 (12.7) IFA Psaroulaki et al., 2010
Brown rat Rattus norvegicus http://maps.iucnredlist.org/map.html?id=19353
Czech Rep. 41/286 (14.3) CFT Syrucek and Raska, 1956
n.a. ISO Syrucek and Raska, 1956
Germany 7/524 (1.3) PCR Runge et al., 2013
India 2/21 (9.5) ISO Yadav and Sethi, 1980
3/21 (14.3) CAT Yadav and Sethi, 1980
75
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Rodentia Muridae
Brown rat Rattus norvegicus http://maps.iucnredlist.org/map.html?id=19353
Japan 0/54 (0.0) ELISA Ejercito et al., 1993
Netherl. 23/146 (15.7) ELISA Reusken et al., 2011
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
House mouse Mus musculus http://maps.iucnredlist.org/map.html?id=13972
Czech Rep.
1/78 (1.3) MAT Rehacek et al., 1977
14/155 (9) CFT Syrucek and Raska, 1956
n.a. ISO Proreshnaya et al., 1960
n.a. ISO Syrucek and Raska, 1956
F. USSR 1/6 (16.6) CFT Yevdoshenko and Proreshnaya, 1961
n.a. ISO Yevdoshenko and Proreshnaya, 1961
Fr. Guiana 0/58 (0.0) IFA Gardon et al., 2001
India 0/4 (0.0) CAT Yadav and Sethi, 1980
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
Tamarisk gerbil Meriones tamariscinus http://maps.iucnredlist.org/map.html?id=13169 F. USSR 0/16 (0.0) CFT Yevdoshenko and Proreshnaya, 1961
n.a. No ISO Yevdoshenko and Proreshnaya, 1961
Long-tailed field mouse
Apodemus sylvaticus http://maps.iucnredlist.org/map.html?id=1904
Czech Rep. n.a. No ISO Syrucek and Raska, 1956
Italy 2/101 (19.8) PCR Pascucci et al., 2015
Slovakia 0/3 (0.0) PCR Smetanova et al., 2006
76
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Rodentia
Muridae
Long-tailed field mouse
Apodemus sylvaticus http://maps.iucnredlist.org/map.html?id=1904 Spain 1/162 (0.6) PCR Barandika et al., 2008
UK 48/307 (15.6) ELISA Meredith et al., 2014
Yellow-necked field mouse
Apodemus flavicollis http://maps.iucnredlist.org/map.html?id=1892
Czech Rep. 1/202 (0.5) MAT Rehacek et al., 1977
11/212 (5.2) CFT Syrucek and Raska, 1956
Slovakia 1/38 (2.6) PCR Smetanova et al., 2006
Spain 0/3 (0.0) PCR Barandika et al., 2008
Barbary striped grass mouse
Lemniscomys barbarus http://maps.iucnredlist.org/map.html?id=11487 Kenya n.a. ISO Heisch, 1960
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
Sciuridae
Yellow-pine chipmunk Tamias amoenus http://maps.iucnredlist.org/map.html?id=42569 USA n.a. ISO Burgdorfer et al., 1963
1/85 (1.2) MAT Rieman et al., 1979
California ground squirrel
Spermophilus beecheyi http://maps.iucnredlist.org/map.html?id=20481 USA
19/145 (13.1) CFT Enright et al., 1971
6/26 (23.0) CFT Enright et al., 1969
7/112 (6.2) MAT Rieman et al., 1979
n.a. ISO Enright et al., 1971
Arizona black-tailed prairie dog
Cynomys ludovicianus http://maps.iucnredlist.org/map.html?id=6091 USA n.a. CFT Enright et al., 1971
Cliff chipmunk Tamias dorsalis http://maps.iucnredlist.org/map.html?id=42571 USA 0/2 (0.0) CFT Stoenner et al., 1960
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 ground squirrel
Spermophilus lateralis http://maps.iucnredlist.org/map.html?id=42468 USA 0/34 (0.0) MAT Rieman et al., 1979
n.a. ISO Burgdorfer et al., 1963
Eastern gray squirrel Sciurus carolinensis http://maps.iucnredlist.org/map.html?id=42462 USA 4/72 (5.5) CFT Enright et al., 1971
n.a. No ISO Enright et al., 1971
New Mexico least chipmunk
Tamias minimus http://maps.iucnredlist.org/map.html?id=42572 1/8 (12.5) CFT Stoenner et al., 1960
77
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Rodentia Sciuridae
Long-clawed ground squirrel
Spermophilopsis leptodactylus
http://maps.iucnredlist.org/map.html?id=20471 Former USSR
n.a. ISO Zhmaeva et al., 1955
Red squirrel Tamiasciurus hudsonicus
http://maps.iucnredlist.org/map.html?id=42587 Canada n.a. (40.0) PCR Thompson et al., 2012
Carolina flying squirrel
Glaucomys sabrinus http://maps.iucnredlist.org/map.html?id=39553 Canada n.a. (38.0) PCR Thompson et al., 2012
Eurasian red squirrel Sciurus vulgaris http://maps.iucnredlist.org/map.html?id=20025 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956
Tien Shan ground squirrel
Spermophilus relictus http://maps.iucnredlist.org/map.html?id=20491
Czech Rep. n.a. ISO Proreshnaya et al., 1960
Former USSR
12/182 (6.6) CFT Yevdoshenko and Proreshnaya, 1961
Czech Rep. n.a. ISO Yevdoshenko and Proreshnaya, 1961
Rock squirrel Spermophilus variegatus http://maps.iucnredlist.org/map.html?id=20495 USA 0/1 (0%) CFT Stoenner et al., 1960
Sonoma chipmunk Tamias sonomae http://maps.iucnredlist.org/map.html?id=42581 USA 2/9 (22.2%) CFT Enright et al., 1971
n.a. No ISO Enright et al., 1971
Southern flying squirrel
Glaucomys volans http://maps.iucnredlist.org/map.html?id=9240 Canada n.a. (65.0) PCR Thompson et al., 2012
Townsend's ground squirrel
Spermophilus townsendii
http://maps.iucnredlist.org/map.html?id=20476 USA 0/1 (0.0) CFT Stoenner et al., 1960
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 squirrel
Ammospermophilus leucurus
http://maps.iucnredlist.org/map.html?id=42452 USA 0/104 (0.0) CFT Stoenner et al., 1960
Soricomorpha Soricidae
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-toothed shrew
Crocidura leucodon http://maps.iucnredlist.org/map.html?id=29651 Czech Rep. 0/2 (0.0) MAT Rehacek et al., 1977
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
78
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Accipitriformes Accipitridae
Black kite Milvus migrans http://maps.iucnredlist.org/map.html?id=22734972 Spain 1/7 (14.0) PCR Astobiza et al., 2011
Bonelli's eagle Aquila fasciata http://maps.iucnredlist.org/map.html?id=22696076 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014*
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
Eurasian buzzard Buteo buteo http://maps.iucnredlist.org/map.html?id=61695117 Cyprus 1/2 (50.0) PCR Psaroulaki et al., 2014
1/4 (25.0) PCR Ioannou et al., 2009
Eurasian griffon vulture
Gyps fulvus http://maps.iucnredlist.org/map.html?id=22695219 Portugal 0/19 (0.0) PCR Cumbassá et al., 2015
Spain 1/9 (11.0) PCR Astobiza et al., 2011
Eurasian sparrowhawk
Accipiter nisus http://maps.iucnredlist.org/map.html?id=22695624 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014
0/2 (50.0) PCR Ioannou et al., 2009
European honey buzzard
Pernis apivorus http://maps.iucnredlist.org/map.html?id=22694989 Cyprus 1/3 (33.3) PCR Ioannou et al., 2009
4/4 (100.0) PCR Psaroulaki et al., 2014
Hen harrier Circus cyaneus http://maps.iucnredlist.org/map.html?id=22727733 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014
2/3 (66.6) PCR Ioannou et al., 2009
Long-legged buzzard Buteo rufinus http://maps.iucnredlist.org/map.html?id=22736562 Cyprus
1/2 (50.0) PCR Psaroulaki et al., 2014
1/3 (66.6) PCR Ioannou et al., 2009
India n.a. (0.0) CAT Yadav and Sethi, 1980
Northern goshawk Accipiter gentilis http://maps.iucnredlist.org/map.html?id=22695683 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014
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
Anseriformes Anatidae
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
Common teal Anas crecca http://maps.iucnredlist.org/map.html?id=22729717 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
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
Mallard Anas platyrhynchos http://maps.iucnredlist.org/map.html?id=22680186
Cyprus 1/1 (100.0) PCR Ioannou et al., 2009
1/2 (50.0) PCR Psaroulaki et al., 2014
India n.a. (0.0) CAT Yadav and Sethi, 1980
Japan n.a. (0.0) PCR To et al., 1998
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
Whooper swan Cygnus cygnus http://maps.iucnredlist.org/map.html?id=22679856 Japan n.a. (0.0) PCR To et al., 1998
0/10 (0.0) MAT To et al., 1998
Bucerotiformes Upupidae Common hoopoe Upupa epops http://maps.iucnredlist.org/map.html?id=22682655 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
79
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
Charadriiformes
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
Burhinidae Eurasian thick-knee Burhinus oedicnemus http://maps.iucnredlist.org/map.html?id=45111439 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
Laridae
Black-headed gull Larus ridibundus http://maps.iucnredlist.org/map.html?id=22694420 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/2 (0.0) PCR Psaroulaki et al., 2014
Caspian gull Larus cachinnans http://maps.iucnredlist.org/map.html?id=22735929 Cyprus 1/4 (25.0) PCR Ioannou et al., 2009
2/3 (66.6) PCR Psaroulaki et al., 2014
Lesser black-backed gull
Larus fuscus http://maps.iucnredlist.org/map.html?id=22694373 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014
1/2 (50.0) 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 Ioannou et al., 2009
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
Ciconiiformes Ciconiidae White stork Ciconia ciconia http://maps.iucnredlist.org/map.html?id=22697691 Cyprus
1/1 (100.0) PCR Ioannou et al., 2009
1/1 (100.0) PCR Psaroulaki et al., 2014
Spain 0/5 (0.0) PCR Astobiza et al., 2011
Columbiformes Columbidae
Band-tailed pigeon Patagioenas fasciata http://maps.iucnredlist.org/map.html?id=22725264 USA 3/31 (9.6) MAT Rieman et al., 1979
Common woodpigeon Columba palumbus http://maps.iucnredlist.org/map.html?id=22690103 Cyprus 3/10 (30.0) PCR Ioannou et al., 2009
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
Mourning dove Zenaida macroura http://maps.iucnredlist.org/map.html?id=22690736 USA 0/9 (0.0) MAT Rieman et al., 1979
Rock dove Columba livia http://maps.iucnredlist.org/map.html?id=22690066
India 1/11 (9.0) CAT Yadav and Sethi, 1980
Iran n.a. (7.9) n.a. Mostafavi et al., 2012
France n.a. ISO Stein et al., 1999
Japan n.a. (0.0) PCR To et al., 1998
4/100 (4.0) MAT To et al., 1998
Coraciiformes
Alcedinidae Common kingfisher Alcedo atthis http://maps.iucnredlist.org/map.html?id=22683027 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014
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 Ioannou et al., 2009
0/2 (0.0) PCR Psaroulaki et al., 2014
80
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Falcnoniformes
Acciprinidae Indian vulture Gyps indicus http://maps.iucnredlist.org/map.html?id=22729731 India n.a. (0.0) CAT Yadav and Sethi, 1980
Falconidae
n.a. n.a. Spain 0/15 (0.0) PCR Astobiza et al., 2011
Common kestrel Falco tinnunculus http://maps.iucnredlist.org/map.html?id=22696362 Cyprus
4/11 (36.3) PCR Ioannou et al., 2009
9/19 (47.3) PCR Psaroulaki et al., 2014
Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956
Eleonora's falcon Falco eleonorae http://maps.iucnredlist.org/map.html?id=22696442 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
Eurasian hobby Falco subbuteo http://maps.iucnredlist.org/map.html?id=22696460 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
Peregrine falcon Falco peregrinus http://maps.iucnredlist.org/map.html?id=45354964 Cyprus 1/1 (100.0) PCR Psaroulaki et al., 2014
2/4 (50.0) PCR Ioannou et al., 2009
Red-footed falcon Falco vespertinus http://maps.iucnredlist.org/map.html?id=22696432 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/2 (0.0) PCR Psaroulaki et al., 2014
Saker falcon Falco cherrug http://maps.iucnredlist.org/map.html?id=22696495 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
Galliformes Phasianidae
Black francolin Francolinus francolinus http://maps.iucnredlist.org/map.html?id=22678719 Cyprus 0/2 (0.0) PCR Ioannou et al., 2009
0/2 (0.0) PCR Psaroulaki et al., 2014
Chukar Alectoris chukar http://maps.iucnredlist.org/map.html?id=22678691 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014
0/2 (0.0) PCR Ioannou et al., 2009
Common quail Coturnix coturnix http://maps.iucnredlist.org/map.html?id=22678944 Cyprus 0/3 (0.0) PCR Ioannou et al., 2009
1/2 (50.0) PCR Psaroulaki et al., 2014
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
Common pheasant Phasianus colchicus http://maps.iucnredlist.org/map.html?id=45100023 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956
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
Gruiformes
Gruidae Common crane Grus grus http://maps.iucnredlist.org/map.html?id=22692146 Cyprus 1/1 (100.0) PCR Ioannou et al., 2009
1/1 (100.0) PCR Psaroulaki et al., 2014
Rallidae
American coot Fulica americana http://maps.iucnredlist.org/map.html?id=62169677 USA 15/33 (45.4) MAT Rieman et al., 1979
Common coot Fulica atra http://maps.iucnredlist.org/map.html?id=22692913 Cyprus 1/5 (20.0) PCR Ioannou et al., 2009
3/4 (75.0) PCR Psaroulaki et al., 2014
Common moorhen Gallinula chloropus http://maps.iucnredlist.org/map.html?id=62120190 Cyprus 1/5 (20.0) PCR Ioannou et al., 2009
5/7 (71.4) PCR Psaroulaki et al., 2014
Corncrake Crex crex http://maps.iucnredlist.org/map.html?id=22692543 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014
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
81
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Passeriformes
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
Corvidae
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
Carrion crow Corvus corone http://maps.iucnredlist.org/map.html?id=22706016
Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956
India n.a. ISO Yadav and Sethi, 1980
Japan 13/173 (7.5.0) PCR To et al., 1998
64/173 (37.0) MAT To et al., 1998
Spain 0/6 (0.0) PCR Astobiza et al., 2011
Common raven Corvus corax http://maps.iucnredlist.org/map.html?id=22706068 Poland n.a. ISO Tylewska-Wierzbanowska et al., 1991
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
Jungle crow Corvus macrorhynchos http://maps.iucnredlist.org/map.html?id=22706019 Japan 5/41 (12.2) PCR To et al., 1998
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
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
Emberizidae
Canyon towhee Melozone fuscus http://maps.iucnredlist.org/map.html?id=22721331 USA 0/4 (0.0) MAT Rieman et al., 1979
Golden-crowned sparrow
Zonotrichia atricapilla http://maps.iucnredlist.org/map.html?id=22721091 USA 8/10 (80.0) MAT Rieman et al., 1979
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 sparrow
Zonotrichia leucophrys http://maps.iucnredlist.org/map.html?id=22721088 USA 34/48 (70.8) MAT Rieman et al., 1979
Yellowhammer Emberiza citrinella http://maps.iucnredlist.org/map.html?id=22720878 Czech Rep. 1/29 (3.4) CFT Syrucek and Raska, 1956
Fringillidae
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
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
82
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Passeriformes
Hirundinidae
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
Swallow n.a. India 1/200 (0.5) ISO Yadav and Sethi, 1980
India 6/200 (3.0) CAT Yadav and Sethi, 1980
Icteridae Brewer´s blackbird Euphagus
cyanocephalus http://maps.iucnredlist.org/map.html?id=22724332 USA 6/18 (33.3) MAT Rieman et al., 1979
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
Laniidae Lesser grey shrike Lanius minor http://maps.iucnredlist.org/map.html?id=22705038 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009 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
Motacillidae Grey wagtail Motacilla cinerea http://maps.iucnredlist.org/map.html?id=22718392 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956 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
Muscicapidae
Black redstart Phoenicurus ochruros http://maps.iucnredlist.org/map.html?id=22710051 Czech Rep. 2/22 (9.0) CFT Syrucek and Raska, 1956 Common redstart Phoenicurus phoenicurus http://maps.iucnredlist.org/map.html?id=22710055 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956 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
Paridae
Black-capped chickadee
Parus atricapillus http://maps.iucnredlist.org/map.html?id=22711716 Czech Rep. 0/2 (0.0) CFT Syrucek and Raska, 1956
Blue tit Parus caeruleus http://maps.iucnredlist.org/map.html?id=22711944 Czech Rep. 0/3 (0.0) CFT Syrucek and Raska, 1956
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
Passeridae
Eurasian tree sparrow Passer montanus http://maps.iucnredlist.org/map.html?id=22718270 Czech Rep. 0/8 (0.0) CFT Syrucek and Raska, 1956
House sparrow Passer domesticus http://maps.iucnredlist.org/map.html?id=22718174 Czech Rep. 5/31 (16.1) CFT Syrucek and Raska, 1956
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
Sturnidae Common myna Acridotheres tristis http://maps.iucnredlist.org/map.html?id=22710921 India 19/69 (27.5) CAT Yadav and Sethi, 1980
3/69 (4.3) ISO Yadav and Sethi, 1980
83
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Passeriformes
Sturnidae
n.a. n.a. Spain 0/4 (0.0) PCR Astobiza et al., 2011
Common starling Sturnus vulgaris http://maps.iucnredlist.org/map.html?id=22710886 Czech Rep. 0/10 (0.0) CFT Syrucek and Raska, 1956
USA 18/157 (11.4) MAT Rieman et al., 1979
Sylviidae
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
n.a. n.a. Spain 0/1 (0.0) PCR Astobiza et al., 2011
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
Turdidae
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
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
Pelecaniformes Ardeidae
Black-crowned night-heron
Nycticorax nycticorax http://maps.iucnredlist.org/map.html?id=22697211 Cyprus 1/1 (100.0) PCR Ioannou et al., 2009
1/1 (100.0) PCR Psaroulaki et al., 2014
Common little bittern Ixobrychus minutus http://maps.iucnredlist.org/map.html?id=22735766 Cyprus 1/1 (100.0) PCR Ioannou et al., 2009
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
Grey heron Ardea cinerea http://maps.iucnredlist.org/map.html?id=22696993 Cyprus 1/2 (50.0) PCR Psaroulaki et al., 2014
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
Little egret Egretta garzetta http://maps.iucnredlist.org/map.html?id=62774969 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
n.a. n.a. Spain 0/12 (0.0) PCR Astobiza et al., 2011
Purple heron Ardea purpurea http://maps.iucnredlist.org/map.html?id=22697031 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
Phoenicopteriformes Phoenicopteridae American flamingo Phoenicopterus ruber http://maps.iucnredlist.org/map.html?id=22729706 Cyprus 1/2 (50.0) PCR Psaroulaki et al., 2014
1/3 (33.3) PCR Ioannou et al., 2009
Piciformes Picidae
Great spotted woodpecker
Dendrocopos major http://maps.iucnredlist.org/map.html?id=22681124 Czech Rep. 0/7 (0.0) CFT Syrucek and Raska, 1956
Grey-faced woodpecker
Picus canus http://maps.iucnredlist.org/map.html?id=22726503 Czech Rep. 0/1 (0.0) CFT Syrucek and Raska, 1956
Podicipediformes Podicipedidae
Great crested grebe Podiceps cristatus http://maps.iucnredlist.org/map.html?id=22696602 Cyprus 0/1 (0.0) PCR Ioannou et al., 2009
0/1 (0.0) PCR Psaroulaki et al., 2014
Little grebe Tachybaptus ruficollis http://maps.iucnredlist.org/map.html?id=22696545 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014
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
84
Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Psittaciformes Psittacidae Rose-ringed parakeet Psittacula krameri http://maps.iucnredlist.org/map.html?id=22685441 India 1/56 (1.8) ISO Yadav and Sethi 1980
13/56 (23.2) CAT Yadav and Sethi 1980
Strigiformes
Strigidae
Great horned owl Bubo virginianus http://maps.iucnredlist.org/map.html?id=61752071 USA 0/1 (0.0) MAT Rieman et al., 1979
Little owl Athene noctua http://maps.iucnredlist.org/map.html?id=22689328 Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014
0/2 (0.0) PCR Ioannou et al., 2009
n.a. n.a. Spain 0/14 (0.0) PCR Astobiza et al., 2011
Northern long-eared owl
Asio otus http://maps.iucnredlist.org/map.html?id=22689507 Cyprus 1/4 (25.0) PCR Ioannou et al., 2009
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
Tytonidae Common barn owl Tyto alba http://maps.iucnredlist.org/map.html?id=22688504 Cyprus
2/5 (40.0) PCR Ioannou et al., 2009
3/5 (60.0) PCR Psaroulaki et al., 2014
n.a. n.a. Spain 0/23 (0.0) PCR Astobiza et al., 2011
Suliformes Phalacrocoracidae
Great cormorant Phalacrocorax carbo http://maps.iucnredlist.org/map.html?id=22696792 Cyprus 0/3 (0.0) PCR Ioannou et al., 2009
0/5 (0.0) PCR Psaroulaki et al., 2014
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
85
Clase Order Family Common name Scientific name Distribution Country Pos/N (Prev.) Method Reference
Amphibia Anura
Bufonidae Cane toad Rhinella marina http://maps.iucnredlist.org/map.html?id=41065 Fr. Guiana 0/21 (0.0) ELISA Gardon et al., 2001
Toads Bufo spp. India 0/66 (0.0) CAT Yadav and Sethi, 1979
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
Sauropsida
Squamata
Natricidae Snakes Natrix natrix & Naja naja http://maps.iucnredlist.org/map.html?id=14368 India 11/20 (55.0) PCR Yadav and Sethi, 1979
11/48 (23.0) CAT Yadav and Sethi, 1979
Colubridae Chinese rat snake Ptyas korros http://www.catalogueoflife.org/col/details/species India
1/2 (50.0) PCR Yadav and Sethi, 1979
2/5 (40.0) CAT Yadav and Sethi, 1979
7/23 (30.4) CAT Yadav and Sethi, 1980
n.a. ISO Yadav and Sethi, 1980
Varanidae Indian monitor Varanus indicus http://maps.iucnredlist.org/map.html?id=178416
India
1/1 (100.0) PCR Yadav and Sethi, 1979
n.a. (0.0) CAT Yadav and Sethi, 1980
Testudines Geoemydidae Tortoise
Pangshura tecta http://maps.iucnredlist.org/map.html?id=46370 India n.a. (0.0) CAT Yadav and Sethi, 1980
Kachuga spp. http://maps.iucnredlist.org/map.html?id=46370 India 2/16 (12.5) CAT Yadav and Sethi, 1979
4/7 (57.1) PCR Yadav and Sethi, 1979
Actinopterygii
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
Cypriniformes Cyprinidae Mrigal Cirrhinus mrigala http://maps.iucnredlist.org/map.html?id=166146 India 0/15 (0.0) CAT Yadav and Sethi, 1979
2/2 (100.0) PCR Yadav and Sethi, 1979
86
Scientific Name Host Host scientific name Country Pos/N (Prev) Method Reference
Amblyomma americanum Questing USA n.a. (+) PCR Klyachko et al., 2007
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
Amblyomma latum Python Python sp. Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009
Questing Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009
Amblyomma triguttatum
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
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
Dermacentor marginatus
Questing
Slovakia ISO Rehacek et al., 1991
0/16 (0) PCR Smetanova et al., 2006
Slovakia & Hungary 2/43 (4.6) PCR Spitalska et al., 2003
Spain 18/265 (27.7) PCR/RLB Toledo et al., 2009
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, wild boar, red fox, European hedgehog, beech marten)
Cervus elaphus, Sus scrofa, Vulpes vulpes, Erinaceus europaeus, Martes foina
Spain 0/23 (0.0) PCR/RLB Toledo et al., 2009
Dermacentor reticulatus
European badger Meles meles Spain 0/3 (0.0) PCR Astobiza et al., 2011
Questing Slovakia
ISO Rehacek et al., 1991
0/9 (0.0) PCR Smetanova et al., 2006
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
87
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
Haemaphysalis concinna Questing
Spain n.a. (0.0) PCR Barandika et al., 2008
Slovakia n.a. (+) ISO Rehacek et al., 1991
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
Haemaphysalis hispanica European hare & rabbit
Lepus europaeus, Oryctolagus cuniculus
Spain 0/5 (0.0) PCR/RLB Toledo et al., 2009
Questing Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009
Haemaphysalis humerosa Common northern bandicoot
Isoodon macrourus Australia 0/250 (0.0) PCR Cooper et al., 2013
Haemaphysalis inermis
Questing Slovakia n.a. (+) ISO Rehacek et al., 1991
Slovakia & Hungary 0/7 (0.0) PCR Spitalska et al., 2003
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
Haemaphysalis punctata
Cypriot mouflon Ovis orientalis ophion Cyprus 2/2 (100.0) PCR Ioannou et al., 2011*
2/2 (100.0) PCR Psaroulaki et al., 2014a
Questing
Slovakia n.a. (+) ISO Rehacek et al., 1991
Spain n.a. (+) PCR Barandika et al., 2008
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
Haemaphysalis sulcata Cypriot mouflon Ovis orientalis ophion Cyprus 5/41 (12.2) PCR Ioannou et al., 2011
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
88
Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference
Hyalomma excavatum Cypriot mouflon Ovis orientalis ophion Cyprus 2/15 (13.3) PCR Ioannou et al., 2011
2/15 (13.3) PCR Psaroulaki et al., 2014a
Hyalomma lusitanicum
Birds (Booted eagle, common buzzard, red kite)
Hieraaetus pennatus, Buteo buteo, Milvus milvus
Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009
Questing Spain 61/701 (8.7) PCR/RLB Toledo et al., 2009
Red deer Cervus elaphus Spain 3/10 (30.0) PCR/RLB Toledo et al., 2009
Wild mammals (wild boar, red fox, European hedgehog, beech marten)
Sus scrofa, Vulpes vulpes, Erinaceus europaeus, Martes foina
Spain 0/11 (0.0) PCR/RLB Toledo et al., 2009
Hyalomma marginatum
Birds (Booted eagle, common buzzard, red kite)
Hieraaetus pennatus, Buteo buteo, Milvus milvus
Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009
Common blackbird, blackcap, common Redstart, whinchat, whitethroat, nightingale, European honey buzzard
Turdus merula, Sylvia atricapilla, Phoenicurus phoenicurus, Saxicola rubetra, Sylvia communis, Luscinia megarhynchos, Pernis apivorus
Italy 10/37 (27.0) PCR Toma et al., 2014
Cypriot mouflon Ovis orientalis ophion Cyprus 0/1 (0.0) PCR Ioannou et al., 2011
0/1 (0.0) PCR Psaroulaki et al., 2014a
Hyalomma rufipes
Common blackbird, blackcap, common Redstart, whinchat, whitethroat, nightingale, European honey buzzard
Turdus merula, Sylvia atricapilla, Phoenicurus phoenicurus, Saxicola rubetra, Sylvia communis, Luscinia megarhynchos, Pernis apivorus
Italy 29/71 (40.0) PCR Toma et al., 2014
Hyalomma spp.
Eurasian reed warbler Acrocephalus scirpaceus France 0/1 (0.0) PCR Socolovschi et al., 2012
Common blackbird, blackcap, common Redstart, whinchat, whitethroat, nightingale, European honey buzzard
Turdus merula, Sylvia atricapilla, Phoenicurus phoenicurus, Saxicola rubetra, Sylvia communis, Luscinia megarhynchos, Pernis apivorus
Italy 29/71 (40.0) PCR Toma et al., 2014
89
Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference
Ixodes acuminatus Bank vole Myodes glareolus Italy 2/88 (2.2) PCR Pascucci et al., 2015*
Wood mouse Apodemus sylvaticus Italy 4/88 (4.5) PCR Pascucci et al., 2015*
Ixodes canisuga European badger Meles meles Spain 0/18 (0.0) PCR Astobiza et al., 2011
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
Ixodes gibossus
Cypriot mouflon Ovis orientalis ophion Cyprus 0/3 (0.0) PCR Ioannou et al., 2011
0/3 (0.0) PCR Psaroulaki et al., 2014a
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
Ixodes hexagonus
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
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
Ixodes holocyclus Common northern bandicoot
Isoodon macrourus Australia 10/30 (33.3) PCR Cooper et al., 2013
Ixodes ricinus
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
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
90
Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference
Ixodes ricinus
Questing
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
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
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
Red deer Cervus elaphus Spain 0/40 (0.0) PCR Astobiza et al., 2011
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
Ixodes spp.
Bank vole Myodes glareolus Italy n.a. (0.0) PCR Pascucci et al., 2015
Common blackbird, blackcap, common redstart, whinchat, whitethroat, nightingale, European honey buzzard
Turdus merula, Sylvia atricapilla, Phoenicurus phoenicurus, Saxicola rubetra, Sylvia communis, Luscinia megarhynchos, Pernis apivorus
Italy 1/6 (16.6) PCR Toma et al., 2014
91
Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference
Ixodes spp.
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
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 guillemot
Fratercula cirrhata, Uria lomvia
USA 0/20 (0.0) PCR Duron et al., 2014
Ixodes ventalloi
Chukar partridge Alectoris chukar Cyprus 3/15 (20.0) PCR Ioannou et al., 2009
Cyprus 2/2 (100.0) PCR Psaroulaki et al., 2014a
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
Rhipicephalus bursa
Cypriot mouflon Ovis orientalis ophion Cyprus 14/35 (40.0) PCR Ioannou et al., 2011
14/35 (40.0) PCR Psaroulaki et al., 2014a
Questing Spain n.a. (0.0) PCR Barandika et al., 2008
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
Rhipicephalus pusillus
Birds H. pennatus, B. buteo, M. milvus
Spain 0/1 (0.0) PCR/RLB Toledo et al., 2009
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
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
Wild mammals C. elaphus, S. scrofa, V. vulpes, E. europaeus, M. foina
Spain 0/28 (0.0) PCR/RLB Toledo et al., 2009
92
Scientific Name Host Name scientific host Country Pos./N (Prev.) Method Reference
Rhipicephalus sanguineus
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
Questing Spain n.a. (0.0) PCR Barandika et al., 2008
Red fox Vulpes vulpes Spain 2/14 (14.3) PCR/RLB Toledo et al., 2009
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, wild boar, European hedgehog, beech marten)
C. elaphus, S. scrofa, E. europaeus, M.es foina
Spain 0/24 (0.0) PCR/RLB Toledo et al., 2009
Rhipicephalus turanicus
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
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
93
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
Ornithodoros capensis s.l.
Cape Verde shearwater, brown booby
Calonectris edwardsii, Sula leucogaster
Cape Verde 16/16 (100.0) PCR Duron et al., 2014
Humboldt penguin Spheniscus humboldti Chile 3/3 (100.0) PCR Duron et al., 2014
Peruvian pelican, Peruvian booby
Pelecanus thagus, Sula variegata
Peru 5/5 (100.0) PCR Duron et al., 2014
Sooty tern Onychoprion fuscatus Mozambique 28/28 (100.0) PCR Duron et al., 2014
Yellow-legged gull Larus michahellis Spain 20/20 (100.0) PCR Duron et al., 2014
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
Ctenocephalides felis
European hare Lepus europaeus Cyprus 0/4 (0.0) PCR Psaroulaki et al., 2014b
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
Xenopsylla cheopis European hare Lepus europaeus Cyprus 0/1 (0.0) PCR Psaroulaki et al., 2014b
Rats Rattus sp. Cyprus 9/83 (10.3) PCR Psaroulaki et al., 2014b
94
Order Family Common name Scientific name Country Origin Pos/N (Prev.) Method Reference
Artiodactyla Bovidae
Arabian Oryx Oryx leucoryx
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
USA C 0/8 (0.0) CFT Greth at al., 1992
Qatar C 1/2 (50.0) CFT Greth at al., 1992
Saudi Arabia C 18/189 (9.5) CFT Greth at al., 1992
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
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
Mountain gazelle Gazella gazella Arab Emirates C 0/3 (0.0) ELISA Chaver et al., 2012
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
Sand gazelle Gazella leptoceros Arab Emirates C 2/6 (33.3) ELISA Chaver et al., 2012
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
95
Order Family Common name Scientific name Country Origin Pos/N (Prev.) Method Reference
Artiodactyla
Bovidae Water buffalo Bubalus bubalis Italy F
12/1012 (1.2) SERO Galiero et al., 1996
14/164 (17.5) PCR Perugini et al., 2009
White antelope Addax nasomaculatus Portugal Z 2/15 (13.3) PCR Cumbassá et al., 2015
Cervidae
Fallow deer Dama dama Slovakia Z 20/60 (33.3) ELISA Dorko et al., 2009
Germany F + CFT/ELISA Simmert et al., 1998
Sika deer Cervus nippon Korea C 0/30 (0.0) ELISA Jang et al., 2011
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
Red deer Cervus elaphus
Portugal Z 0/1 (0.0) PCR Cumbassá et al., 2015
Spain
F 6/23 (26.1) PCR González-Barrio et al., 2015
F 9/25 (36.0) ELISA González-Barrio et al., 2015
F 1/29 (3.4) PCR González-Barrio et al., 2015
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
Wapiti Cervus canadensis Germany Z + n.a. Gaukler and Krauss, 1974
Korea F 10/604 (1.7) ELISA Jang et al., 2011
Carnivora Felidae Lion Panthera leo Italy S
1/8 (12.5) PCR Laricchiuta et al., 2009
2/8 (25.0) IFA Laricchiuta et al., 2009
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
96
Order Family Common name Scientific name Country Origin Pos/N (Prev.) Method Reference
Columbiformes Columbidae Rock dove Columba livia Slovakia SD
2/97 (2.0) MAT Kocianova et al., 1993
12/67 (17.9) MAT Kocianova et al., 1993
16/97 (16.5) MAT Kocianova et al., 1993
Galliformes Phasianidae Common quail Coturnix coturnix Germany E + MAT Schatmz et al., 1977
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
Piciformes Ramphastidae Toucan Ramphastos toco USA C Pathology PCR Shivaprarad et al., 2008
Cacatuidae Cockatiel Nymphicus hollandicus USA C Pathology PCR Shivaprarad et al., 2008
Psittaciformes
Psittacidae Canary-winged parakeet Brotogeris chiriri USA C Pathology PCR Shivaprarad et al., 2008
Psittaculidae
Hawk-headed parrot Deroptyus accipitrinus USA C Pathology PCR Shivaprarad et al., 2008
Red-fronted parakeet Cyanoramphus novaezelandiae USA C Pathology PCR Shivaprarad et al., 2008
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
97
Objetivos
98
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.
99
Capítulo II. Epidemiología
de Coxiella burnetii en
fauna silvestre ibérica
100
Capítulo II. 1
101
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
102
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.
103
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.
104
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
105
(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.
106
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
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).
107
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
108
(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).
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.
aLocation 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.
Table 1. Primers and probe used in the qPCR.
109
(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):
110
Factor Variable
codea
Variable description (measure unit)
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)
SFd Density of sheep farms in the municipality (farms/Km2)
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
(farms/Km2)
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) Densities of domestic ruminants and domestic ruminant farms in the municipality
to which individual deer belong. Domestic ruminant density (densities of cattle
Table 2. Set of variables gathered for risk factor modeling of deer individual exposure to
Coxiella burnetii.
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
111
[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.
(c) Density of humans in the municipality (HUd). Updated human demographic data
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
112
Country or
bioregion
Seroprevalence (%) (Pos/N)b (95%CI)
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)
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.
113
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
114
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.
Sex Age class
Seroprevalence (%)
(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)
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.
115
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.
Data set Variableb Z β SE Pc AIC ΔAIC ED (%)
Unmanaged + Managed +
Farmed
Intercept -2.561 -3.745 1.463 *
364.105 19.206 7.172
HUd 2.707 0.019 0.007 **
Se -2.476 -0.690 0.279 *
AvSpT 2.008 0.204 0.101 *
Rud -1.573 -0.017 0.011 NS
Unmanaged Intercept -1.331 -1932 1.473 NS
220.417 2.525 3.723 HUd 1.788 0.053 0.030 NS
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 **
91.102 16.224 19.572 Se 3.705 1.983 0.535 ***
RuD -3.499 -0.186 0.053 ***
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
Abbreviations of variables are explained in Table 2. c NS, P>0.05; *, P <0.05; **, P <0.01; ***,
P <0.001
116
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
117
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]).
118
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.
119
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
120
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
121
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;
122
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
123
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.
124
Capítulo II. 2
125
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
126
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.
127
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.
128
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
129
(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.
130
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).
131
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
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.
132
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).
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.
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 ¶ § § §
1
*Cbsp, overall seropositivity; CbspLO, seropositivity at location LO; CbspILO, systemic infection (C.
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.
133
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.
134
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.
135
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.
136
Capítulo II. 3
137
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
138
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.
139
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 re-
vaccination every 6 months.
140
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;
141
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
142
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
143
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
144
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
145
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
146
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
147
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
148
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.
149
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.
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
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.
150
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
151
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.
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
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.
152
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.
153
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.
154
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,
155
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
156
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
157
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
158
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.,
159
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
160
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.
161
Capítulo II. 4.
162
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.
163
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.
164
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.
165
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).
166
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.
167
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.
168
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
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.
169
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
170
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).
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
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.
171
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
172
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).
173
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
174
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).
175
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 multi-
host 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.
176
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).
177
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.
178
Su
pp
lem
en
tary
fil
es
Ap
pen
dix
1a.
Min
imu
m s
pan
nin
g t
ree
show
ing t
he
rela
tionsh
ip b
etw
een t
he
obta
ined
ML
VA
gen
oty
pes
id
enti
fied
in t
his
stu
dy (
*)
an
d i
sola
tes
fro
m
div
erse
host
ori
gin
ob
tain
ed f
rom
Co
xiel
la b
urn
etii
2014
coop
erati
ve
data
base
of
ML
VA
Bank
5.0
(htt
p:/
/mlv
a.u
-psu
d.f
r/m
lvav4
/gen
oty
pin
g/i
nd
ex.p
hp
).
Eac
h c
ircl
e re
pre
sents
a u
niq
ue
gen
oty
pe;
the
size
of
the
circ
le c
orr
esp
onds
to t
he
nu
mb
er o
f sa
mp
les
wit
h t
hat
gen
oty
pe
as
sho
wn i
n t
he
leg
end. C
om
ple
te
(17)
and a
lmost
co
mp
lete
(7
) M
LV
A g
enoty
pes
wer
e in
clu
ded
in t
he
analy
sis
179
Ap
pen
dix
1b
. M
inim
um
sp
an
nin
g t
ree
show
ing t
he
rela
tionsh
ip b
etw
een t
he
obta
ined
ML
VA
gen
oty
pes
id
enti
fied
in t
his
stu
dy (
*)
an
d i
sola
tes
fro
m d
iver
se
geo
gra
phic
ori
gin
ob
tain
ed f
rom
Co
xiel
la b
urn
etii
2014
coop
erati
ve
data
bas
e of
ML
VA
Bank
5.0
(htt
p:/
/mlv
a.u
-psu
d.f
r/m
lvav4
/gen
oty
pin
g/i
nd
ex.p
hp
). E
ach c
ircle
repre
sents
a u
niq
ue
gen
oty
pe;
the
size
of
the
circ
le c
orr
esp
on
ds
to t
he
nu
mb
er o
f sa
mp
les
wit
h t
hat
gen
oty
pe
as
show
n i
n t
he
legen
d.
Com
ple
te (
17)
and a
lmost
com
ple
te (
7)
ML
VA
gen
oty
pes
wer
e in
clu
ded
in t
he
analy
sis.
180
Capítulo II. 5.
181
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
182
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
183
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.
184
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.
185
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).
186
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.
187
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
188
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.
189
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,
190
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
191
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.
192
Table 2. Summary of genotypes within genomic groups found in domestic and wild animals analyzed in this study
Host Sample Province Ct
CB
U00
07
CB
U00
71
CB
U01
68
CB
U05
98
CB
U08
81
CB
U18
05
CB
U20
26
CB
U09
52*
Gen
om
ic
gro
up
Goat
(Capra aegagrus hircus) Bulk-tank milk
Sevilla 31,12 IV-
Sevilla 32,48 IV-
Racoon (Procyon lotor)
Spleen
Guadalajara 34,54 I-
Madrid 35,08 I+
Madrid 35,27 I+
Guadalajara 35,19 II-
Guadalajara 35,46 II-
Red deer
(Cervus elaphus)
Milk Cadiz 32,18 I+
Cadiz 34,17 I+
Vaginal swab Cádiz 32,50 I+
Spleen Ciudad Real 33,02 I+
Ciudad Real 33,02 I+
Vaginal swab Cádiz 31,40 VI-
Milk Cádiz 33,22 VII-
Sheep (Ovis aries)
Semen
Ciudad Real 35,29 I-
Ciudad Real 33,63 I+
Ciudad Real 34,23 II-
Ciudad Real 34,46 II-
Ciudad Real 35,29 III+
Ciudad Real 36,53 III+
European wild rabbit
(Oryctolagus cuniculus)
Spleen
Sevilla 35,76 I-
Zaragoza 21,89 I-
Zaragoza 25,12 I-
Vaginal swab Cádiz 28,48 VII-
Cádiz 31.18 VII-
Uterus Cádiz 29.93 VII-
Cádiz 35,46 VII-
Spleen Toledo 32,09 VII-
*: Acute disease antigen A (adaA).
193
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
194
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 &
195
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
196
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).
197
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
198
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
199
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.
200
Capítulo III. Vías de
transmisión de Coxiella
burnetii en fauna silvestre
ibérica.
201
Capítulo III. 1.
202
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
203
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.
204
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.
205
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
206
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.
207
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
208
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.
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
Figure 1. Geographic location of the study hunting estates within peninsular Spain.
209
(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
210
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).
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
Pathogen Shedding routes Collected swabs Primer ref. Sequence (5´-3´) Target products (bp)
ADV Nasal, oral, genital and
milka
Nasal, oral, rectal,
genital
fADVgB1a ATGGCCATCTCGCGGTGC gB gene (334)e
ADVgB1b ACTCGCGGTCCTCCAGCA
fADVgB2a ACGGCACGGGCGTGATC gB gene (195)e
ADVgB2b GGTTCAGGGTACCCCGC
PPV Nasal, oral and genitalb Nasal, oral, rectal,
genital
PVF AGTTAGAATAGGATGCGAGGAA NS1 gene (265)f
PVR AGAGTCTGTTGGTGTATTTATTGG
PCV2 Nasal, oral, rectal and
genitalc
Nasal, oral, rectal,
genital
CVF CGAGAAAGCGAAAGGAACAGA ORF 1 (371)f
CVR GGTAACCATCCCACCACTT
Coxiella burnetii Rectal, genital and
milk in ruminantsd;
unknown in suids
Nasal, oral, rectal,
genital
Trans1 TATGTATCCACCGTAGCCAGTC Transposon-like
repetitive region (687)g Trans2 CCCAACAACACCTCCTTATTC
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.
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
211
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
212
confirmed, perhaps because of the low shedding prevalence of ADV, PPV, PCV2 and C.
burnetii in the studied populations.
Sex Age ADV PPV PCV2 Coxiella burnetii
Male Juvenile 4/7
(57.1; 18.4-90.1)
2/8
(25.0; 3.2-65.1)
3/7
(42.9; 9.9-81.6)
0/8
(0.0; 0.0-37.0)
Sub-adult 8/16
(50.0; 24.7-75.4)
6/16
(37.5; 15.2-64.6)
11/16
(68.8; 41.3-90.0)
0/16
(0.0; 0.0-20.6)
Adult 18/38
(47.4; 31.0-64.2)
28/38
(73.7; 56.9-86.6)
28/38
(73.7; 56.9-86.6)
1/38
(2.6; 0.1-13.8)
Total male 30/61
(49.2; 36.1-62.3)
36/62
(58.1; 44.8-70.5)
42/61
(68.9; 55.7-80.1)
1/62
(1.6; 0.0-8.7)
Female Juvenile 10/12
(83.3; 51.6-97.9)
3/12
(25.0; 5.5-57.2)
8/12
(66.7; 34.9-90.1)
0/12
(0.0; 0.0--26.5)
Sub-adult 7/10
(70.0; 34.8-93.3)
4/10
(40.0; 12.2-73.8)
8/10
(80.0; 44.4-97.5)
0/10
(0.0; 0.0-30.9)
Adult 23/41
(56.1; 39.7-71.5)
29/41
(70.7; 54.5-83.9)
35/41
(85.4; 70.8-94.4)
0/41
(0.0; 0.0-8.6)
Total female 40/63
(63.5; 50.4-75.3)
36/63
(57.1; 44.0-69.6)
51/63
(81.0; 69.1-89.8)
0/63
(0.0; 0.0-5.7)
Total 70/124
(56.0; 47.3-65.3)
72/125
(57.6; 48.4-66.4)
93/124
(75.0; 66.4-82.3)
1/125
(0.8; 0.0-4.4)
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.
Pathogen Sex Swab
Nasal Oral Rectal Genital
ADV Male
0/62
(0.0; 0.0-5.8)
0/62
(0.0; 0.0-5.8)
0/59
(0.0; 0.0-6.1) 5/60
(8.3; 2.8-18.4)
Female
0/54
(0.0; 0.0-6.6) 1/53
(1.9; 0.0-10.1)
1/49
(2.0; 0.0-10.9)
1/50
(2.0; 0.1-10.7)
Total ADV
0/116
(0.0; 0.0-3.1) 1/115
(0.9; 0.0-4.8)
1/108
(0.9; 0.0-5.1)
6/110
(5.5; 2.0-11.5)
PPV Male
3/63
(4.8; 1.0-13.3)
0/62
(0.0; 0.0-5.8) 2/56
(3.6; 0.4-12.3)
1/59
(1.7; 0.0-9.1) Female
1/52
(1.9; 0.0-10.3)
0/49
(0.0; 0.0-7.3) 2/46
(4.3; 0.5-14.8)
0/47
(0.0; 0.0-7.6)
Total PPV
4/115
(3.5; 1.0-8.7)
0/111
(0.0; 0.0-3.2) 4/102
(3.9; 1.1-9.7)
1/106
(0.9; 0.0-5.2)
PCV2 Male
2/62
(3.2; 0.4-11.2)
2/61
(3.3; 0.4-11.4)
0/56
(0.0; 0.0-6.4)
0/59
(0.0; 0.0-6.1)
Female
1/52
(1.9; 0.0-10.3)
1/51
(2.0; 0.0-10.5)
1/47
(2.1; 0.0-11.3)
0/47
(0.0; 0.0-7.6)
Total PCV2
3/114
(2.6; 0.5-7.5)
3/112
(2.7; 0.6-7.6)
1/103
(1.0; 0.0-5.3)
0/106
(0.0; 0.0-3.4)
Coxiella
burnetii
Male
1/62
(1.6; 0.0-8.7)
0/61
(0.0; 0.0-5.9)
0/54
(0.0; 0.0-6.6) 1/58
(1.7; 0.0-9.2) Female
1/50
(1.9; 0.0-10.7)
0/50
(0.0; 0.0-7.1) 1/46
(2.2; 0.0-11.5)
0/47
(0.0; 0.0-7.6)
Total C. burnetii
2/112
(1.8; 0.2-6.3)
0/111
(0.0; 0.0-3.2) 1/100
(1.0; 0.0-5.5)
1/105
(1.0; 0.0-5.2)
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.
213
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
214
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.
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
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.
215
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
216
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
217
(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).
218
Capítulo III. 2.
219
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
220
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
221
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.
222
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.
223
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).
224
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
225
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.
226
Capítulo IV
227
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
228
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
229
el inicio de la vacunación sugiere una possible eficacia de la vacunación en ciervos para
la reducción de la contaminación ambiental por C. burnetii.
230
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.
231
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.
232
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
233
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.
234
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.
235
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
Table 1. Sample size through sample type and time according to the allocation of
animals to vaccinated and unvaccinated groups.
236
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
237
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
238
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
239
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
240
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.
241
Fig
ure
2.
Evolu
tio
n o
f se
rop
revale
nce
(%)
and a
ver
age
anti
body l
evel
s (S
P)
in v
acc
inate
d (
bla
ck l
ine)
and u
nv
acc
inate
d (
gre
y l
ine)
dee
r gro
up
s alo
ng t
he
imp
lem
enta
tion o
f th
e vacc
inati
on e
xp
erim
ent.
Err
or
bar
s re
pre
sent
95%
confi
den
ce
inte
rval
of
sero
pre
vale
nce
and s
tandard
dev
iati
on f
rom
th
e m
en f
or
aver
ag
e
anti
body l
evel
s. R
ed d
ots
dep
ict
the
date
s at
whic
h a
nim
als
wer
e vacc
inate
d.
* S
tati
stic
ally
sig
nif
ican
t dif
fere
nce
s (p
<0.0
5)
bet
wee
n v
acci
nat
ed a
nd u
nvac
cinat
ed g
roups
242
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:
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
243
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.
244
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.
245
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.
246
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
247
(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
248
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.,
249
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.
Fig
ure 4
. B
urd
en o
f C
oxi
ella
burn
etii
sh
ed (
aver
age
qP
CR
Ct
val
ue)
an
d a
ssoci
ated
sta
nd
ard d
evia
tion
(er
ror
bar
s) i
n v
agin
al s
ecre
tion
s, m
ilk a
nd f
aece
s
by
any
coh
ort
of
dee
r st
udie
d a
ccord
ing t
o t
hei
r al
loca
tion
to v
acci
nat
ed (
bla
ck d
iam
on
ds)
an
d u
nvac
cin
ated
(gre
y dia
mon
ds)
gro
ups.
* S
tati
stic
ally
sign
ific
ant
dif
fere
nce
s (p
<0.0
5)
bet
wee
n v
acci
nat
ed a
nd u
nvac
cin
ated
gro
up
s
250
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
251
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
252
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
253
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
254
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
255
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.
256
Capítulo V. Síntesis y
Conclusiones
257
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
258
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
259
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
260
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.
261
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
262
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
263
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
264
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
265
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.
266
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
267
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.
268
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
269
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
270
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
271
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
272
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
273
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
274
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
275
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
276
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.
277
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.
278
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.
279
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.
280
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.
281
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.
282
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
283
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
284
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
285
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
286
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.
287
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
288
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
289
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.
290
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
291
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
292
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 devel- opment 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.
293
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
294
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
295
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
296
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.
297
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]
298
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.
299
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
300
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
301
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,
96
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
302
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.
303
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.
304
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
305
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
306
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
307
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
308
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
309
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
310
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.
311
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
312
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
313
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
314
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.
315
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.
316
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
317
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
318
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
319
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
320
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
321
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.
322
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
323
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
324
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
325
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
326
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.
327
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
328
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
329
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
330
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
331
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
332
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
333
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
334
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
335
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
336
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
337
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
338
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
339
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
340
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.
341
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
342
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
343
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
344
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
