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Risk of Coxiella burnetii transmission via embryo transfer using in vitro early bovine embryos
Theriogenology 81 (2014) 849–853
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Risk of Coxiella burnetii transmission via embryo transfer using in vitro early bovine embryos A. Alsaleh a, F. Fieni a, *, D. Moreno a, E. Rousset b, D. Tainturier a, J.F. Bruyas a, J.L. Pellerin a a
LUNAM University, Oniris, Nantes-Atlantic National College of Veterinary Medicine, Food Science and Engineering, Sanitary Security of Reproduction Biotechnology Unit, Nantes, France b ANSES, Laboratoire de Sophia Antipolis, France
a r t i c l e i n f o
a b s t r a c t
Article history: Received 31 October 2013 Received in revised form 17 December 2013 Accepted 17 December 2013
Coxiella burnetii, an obligate intracellular bacterium of worldwide distribution, is responsible for Q fever. Domestic ruminants are the main source of infection for humans. The objectives of this study were to determine (1) whether C. burnetii would adhere to the intact zona pellucida (ZP-intact) of early in vitro–produced bovine embryos; (2) whether the bacteria would adhere to or infect the embryos (ZP-free) after in vitro infection; and (3) the efficacy of the International Embryo Transfer Society (IETS) washing protocol. One hundred and sixty, eight- to 16-cell bovine embryos produced in vitro, were randomly divided into 16 batches of 10 embryos. Twelve batches (eight ZP-intact and four ZP-free) were incubated in a medium containing C. burnetii CbB1 (Infectiologie Animale et Santé Publique, Institut National de Recherche Agronomique Tours, France). After 18 hours of incubation at 37 C and 5% CO2 in air, the embryos were washed in 10 successive baths of a PBS and 5% fetal calf serum solution in accordance with the IETS guidelines. In parallel, four batches (two ZP-intact and two ZP-free) were subjected to similar procedures but without exposure to C. burnetii to act as controls. Ten washing fluids from each batch were collected and centrifuged for 1 hour at 13,000 g. The embryos and wash pellets were tested using conventional polymerase chain reaction. C. burnetii DNA was found in all ZP-intact and ZPFree embryos after 10 successive washes. It was also detected in the first four washing fluids for ZP-intact embryos and in the 10th wash fluid for two of the four batches of ZPfree embryos. In contrast, none of the embryos or their washing fluids in the control batches were DNA positive. These results demonstrate that C. burnetii adheres to and/or penetrates the early embryonic cells and the ZP of in vitro bovine embryos after in vitro infection, and that the standard washing protocol recommended by the IETS for bovine embryos, failed to remove it. The persistence of these bacteria after washing makes the embryo a potential means of transmission of the bacterium during embryo transfer from infected donor cows to healthy recipients and/or their offspring. Further studies are required to investigate whether enzymatic and/or antibiotic treatment of bovine embryos infected by C. burnetii would eliminate the bacteria from the ZP and to verify if similarly results are obtained with in vivo–derived embryos. Ó 2014 Elsevier Inc. All rights reserved.
Keywords: Coxiella burnetii Embryo transfer International embryo transfer society Zona pellucida Bovine embryo Polymerase chain reaction
1. Introduction
*
Corresponding author. Tel.: þ33 2 40687711; fax: þ33 2 40687748. E-mail address: francis.fi[email protected] (F. Fieni).
0093-691X/$ – see front matter Ó 2014 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.theriogenology.2013.12.015
Embryo transfer (ET) is used to select and diffuse genetic traits without introducing infectious disease agents. Embryos are considered to be free of specific pathogens if their donors or uterine environment are free of the agent or if
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the embryos can resist adherence and infection until the time of collection [1]. Q fever is a zoonotic endemic disease with worldwide distribution caused by an obligate intracellular bacterium, Coxiella burnetii. Domestic ruminants are the main sources for human infection [2,3]. Infection is often asymptomatic in cattle, but it may cause abortion, reproductive failure (metritis, placentitis, and infertility), and economic losses [4–6]. Infection usually occurs via inhalation of contaminated aerosols. Sexual transmission has been implicated in domestic ruminants but has yet to be proven [7,8]. The genital tract, and in particular the placenta, gravid uterus, and mammary tissue, is the preferential target for C. burnetii in pregnant animals [9,10]. Bacterial shedding after abortion has been studied in natural and experimental infections [9,11–15]. Recently, C. burnetii was detected in the genital tract flushing media (oviducts and uterine horns) and tissues of nonpregnant goats with significant bacterial loads [16]. To our knowledge, there are no reported studies on the interaction of bovine embryos and C. burnetii. A recent study conducted in our laboratory (SSBR; Oniris, Nantes, France) demonstrated that in the goat, C. burnetii has a strong tendency to cling to the zona pellucida (ZP) after in vitro infection; the washing procedure recommended by the International Embryo Transfer Society (IETS) for bovine embryos failed to remove it [17]. The IETS has established procedures to reduce the risk of pathogen transmission, which are approved for international exchanges of embryos. However, they must be tested for each pathogen and for embryos from each different species [18]. Most pathogenic agents can be eliminated by washing ZP-intact embryos. However, some pathogens can adhere to the ZP and are not removed by these procedures, resulting in contamination of the embryo and/or uterus; these include Leptospira spp., Brucella ovis, Mycoplasma bovigenitalium, and Mycobacterium avium [19–22]. Today, the detection of Coxiella-DNA in clinical samples using polymerase chain reaction (PCR) has radically changed the diagnosis of Q fever in veterinary medicine. Conventional PCR (C-PCR) and real-time PCR (RT-PCR) techniques are highly sensitive methods; they are also very specific for the detection and quantification of C. burnetii DNA in several biological samples [14,23]. To investigate the risk of C. burnetii transmission via bovine embryo transfer, our study aims to determine whether the embryonic ZP protects early embryo cells against C. burnetii infection, and whether the bacteria adhere to or infect the cells of early bovine embryos (ZPfree) after in vitro infection. We also evaluated the efficacy of the washing procedure recommended by the IETS to decontaminate bovine embryos exposed to C. burnetii in vitro. 2. Materials and methods
intraperitoneal inoculation of three OF1 mice (8 weeks) with 0.2 mL of cow placenta homogenate. The mice were killed 9 days postinoculation and their spleens were sampled and reinoculated into specific pathogen-free embryonated hen eggs. After the third passage in the chicken embryo, Coxiella burnetii was quantified using quantitative-PCR, aliquoted, and frozen at 80 C. This preparation contained 1011 bacteria per mL. To ensure purity, each aliquot used for exposures was diluted with 10 mL PBS then centrifuged twice for 15 minutes at 2000 g; the supernatant was recovered and centrifuged for 1 hour at 13,000 g. The pellet was diluted 1:100 in the exposure medium giving a final calculated concentration of 109 bacteria per mL. 2.2. In vitro production of embryo 2.2.1. In vitro maturation Oocytes were collected by aspiration via an 18-ga needle, from 3 to 8 mm diameter follicles from the ovaries of slaughtered cows. The oocytes were washed four times in HEPES-buffered Tissue Culture Medium 199 (TCM 199, M7528) and 0.4 mg/mL BSA fraction V (A9647) and then matured in 500 mL bicarbonate buffered TCM 199 (M4530) containing 60 IU/mL penicillin, 60 mg/mL streptomycin, and 10% heat-treated calf serum (CVFSVF00-01; Eurobio, Les Ulis, France) for 24 hours at 39 C in an atmosphere of 5% CO2 in humidified air. 2.2.2. In vitro fertilization After maturation, the oocytes were washed in 2 mL of HEPES-buffered TCM 199 (M7528) and 0.4 mg/mL BSA fraction V (A9647), and once in fertilization medium, IVFTyrode’s albumin lactate pyruvate medium supplemented with 6 mg/mL fatty acid-free BSA fraction V (A8806) and 1.7 IU/mL heparin (H3149) before being transferred to fertilization wells. Frozen-thawed bull spermatozoa were used to inseminate the oocytes. Motile spermatozoa were separated on a Percoll discontinuous density gradient (45%–90%) (BoviPure; Nidacon), after centrifugation at 300 g for 20 minutes at room temperature. Spermatozoa were diluted to a final concentration of 4 106 mL. Finally, oocytes and spermatozoa were incubated together for 18 hours at 39 C in a humidified atmosphere of 5% CO2 in air. 2.2.3. In vitro culture The cumulus cells were removed from the oocytes by vortexing them for 2 minutes. Presumptive zygotes were washed in PBS medium before being cultured in groups of approximately 30 embryos in 30-mL droplets of culture medium covered with mineral oil in an atmosphere of 5% O2, 5% CO2, and 90% N2 at 39 C. The culture medium was synthetic oviduct fluid, according to Holm, et al. [24], enriched with 0.4% BSA (A8806). The day of fertilization was considered as Day 0. Embryos were collected on Day-4 postinsemination for in vitro–exposure to CbB1.
2.1. Coxiella burnetii strain 2.3. Removal of the ZP The C. burnetii strain (CbB1) phase I used in this study (provided by IASP, INRA Tours, France) was isolated from the placenta of an aborted cow. It had been purified by
In accordance with the method described by Lamara, et al. [25], the embryos were placed in 1% preincubated
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pronase (Protease, P 6911; Sigma, Paris, France) at 37 C. After 60 to 90 seconds, the embryos were transferred to preincubated acidic Tyrode’s solution (pH 2.1) at 37 C for 90 to 120 seconds. Finally, the embryos were washed three times with minimum essential medium (Sigma) supplemented with 10% fetal calf serum (FCS).
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and a negative control (distilled water) were performed. Samples analyzed for C. burnetii DNA using PCR were considered positive when a band of 337 bp, corresponding to the positive control, was visualized on agarose gel electrophoresis under ultraviolet light. The sensitivity of this PCR method had been proven in our laboratory (SSBR); it detects 10 bacteria per mL of bacterial suspension (data not shown).
2.4. Experimental design 2.6. Real-time PCR procedure Four days after IVF, 160 embryos, were randomly divided into 16 batches of 10 embryos. Twelve batches (eight ZP-intact and four ZP-free) were placed in 1 mL of minimum essential medium (M2414; Sigma) supplemented with 10% FCS, 1% L-glutamine (2 mM final), 1% HEPES (0.01 M final), 2.5 mg/mL1 amphotericin B, and 50 mg/mL gentamicin and containing 109 Coxiella per mL of CbB1 strain (IASP, INRA Tours). After incubation for 18 hours at 37 C in an atmosphere of 5% CO2, the embryos were recovered and washed in batches in 10 successive baths of a PBS and 5% FCS according to the IETS guidelines [18]. A new sterile pipette was used for each successive wash; each wash corresponded to a dilution of 1:100 of the previous medium. In parallel, four batches (two ZP-intact and two ZP-free) were subjected to similar procedures but without exposure to C. burnetii as control group. All the 10 wash baths were collected separately, and centrifuged for 1 hour at 13,000 g. The washed embryos and the pellets of the 10 centrifuged wash baths were frozen at 20 C before examination for evidence of C. burnetii using PCR. 2.5. Conventional PCR procedure DNA was extracted from the batches of embryos and the wash bath pellets using a “QIAamp Blood and body fluid kit, Qiagen, France” in accordance with the manufacturer’s instructions. The detection of Coxiella-DNA was performed by amplifying a DNA fragment (337 bp) located in the transposon-like repetitive region (IS1111) gene, which is present in multiple copies in the Coxiella burnetii genome, using two primers: Trans B: 50 - CAAGAATGATCGTAACGATGCGC-30 (349–371 bp) and Trans M: 50 - CTCGTAATCACCAATCGCTTCG-30 (664–685 bp) (IASP, INRA de Tours). Three microliters of extracted DNA were added to 22 mL of amplification solution. The latter contained 5 mL of ready-to-use solution containing all reagents required for PCR: HOT FIREPol DNA polymerase, Proofreading enzyme, 5 Blend Master Mix Buffer, 7.5 mM MgCl2, 2 mM dNTPs of each, BSA, blue dye, yellow dye, and a compound to increase sample density for direct loading (Solis BioDyne, Estonia), 0.75 mL of both Trans B and Trans M primer (20 mM; Eurofins MWG Operon, Ebersberg, Germany), and 15.5 mL of distilled water DNase–RNase free. Amplification was performed in a thermal cycler (Mastercycler; Eppendorf) on the following program: after initial denaturation at 94 C for 10 minutes, the samples were subjected to a series of 35 cycles of 30 seconds denaturation at 94 C, 1 minute hybridization at 63 C, and a 3-minute elongation phase at 72 C. This was followed by a final elongation phase at 74 C for 10 minutes. Products were visualized by electrophoresis on 1.5% agarose gel. A positive control of C. burnetii (IASP, INRA Tours)
Real-time PCR was used to amplify a DNA fragment of 76 bp from the isocitrate dehydrogenase gene (icd), of which there is only one copy in the C. burnetii genome. The following primers were used: forward, icd-439F: CGT TAT TTT ACG GGT GTG CCA (439–459) and reverse, icd-514R: CAG AAT TTT CGC GGA AAA TCA (494–514), with a TaqMan probe icd-464 TM: FAM-CAT ATT CAC CTT TTC AGG CGT TTT GAC CGT-TAMRA-T (464–492) as described by Klee, et al. [26]. DNase–RNase-free water was used as a negative control. Standard series containing: 2.103, 2.104, 2.105, 2.106, 2.107 C. burnetii per mL (IASP, INRA Tours) were extracted using the QIAamp DNA mini kit (Qiagen) and used as a positive control. Five microliters of extracted DNA were added to 20 mL of RT-PCR reaction mix. The latter was composed of 12.5 mL TaqMan Universal Master Mix II (Applied Biosystems), 2.5 mL of a mixture of forward and reverse primers (0.3 mM; Eurofins MWG Operon), 0.25 mL TaqMan probe (50 nM; Eurofins MWG Operon). Water was added to make a final volume of 20 mL. All RT-PCR reactions were performed in duplicate in an ABIPRISM Sequence Detection System 7300 (Applied Biosystems) as follows: after 2 minutes at 50 C and 10 minutes at 95 C, the samples were subjected to a series of 40 cycles comprising 15 seconds at 95 C and 30 seconds at 60 C. Data were analyzed with the corresponding software. The C. burnetii titers in the samples were calculated in comparison with a standard curve obtained from a standard serial dilution of the bacteria. 2.7. Statistical analysis Fisher’s exact test was used to compare the efficacy of all the 10 embryo-washing procedures in removing Coxiella burnetii from the washing media after in vitro infection. Values of P less than 0.05 were considered to be significant. 3. Results In total, 160 ZP-intact and ZP-free embryos were assigned to 18 batches. Twelve batches were exposed in vitro to Coxiella CbB1 at a concentration of 109 Coxiella per mL; the remaining six batches were used as controls. Coxiella-DNA was detected by C-PCR in all batches of infected ZP-intact and ZP-free embryos (8/8 and 4/4 respectively) after being washed 10 times (Table 1). Bacterial DNA was not detected in any of the embryos in the control batches. For ZP-intact embryo the first three washing media of the infected group were consistently found to be positive (8/8), whereas subsequent washes tested negative (P < 0.05). For ZP-free embryo the first six washing media of the infected group were consistently
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Table 1 Detection of Coxiella burnetii in successive embryo wash baths and batches of infected ZP-intact eight- to 16-cell embryos and infected ZP-free eightto 16-cell embryos after 10 wash cycles, using C-PCR and quantification of Coxiella burnetii in embryo exposure baths and in batches of infected ZPintact eight- to 16-cell embryos or infected ZP-free eight- to 16-cell embryos by RT-PCR. Batch of Exposure embryos bath (Coxiella per mL) (RT-PCR)
Zona pellucida intact 1 2.6 108 2 2.5 108 3 2.4 108 4 2.8 108 5 2.9 108 6 4.4 108 7 4.4 108 8 4.4 108 Zona pellucida free 1 2.9 108 2 2.9 108 3 2.9 108 4 2.9 108
Last positive wash for C. burnetii (C-PCR)
Batch of embryos after 10 wash baths Detection of C burnetii (C-PCR)
Quantification of C burnetii (RT-PCR)
4 3 4 3 3 4 3 5
Positive Positive Positive Positive Positive Positive Positive Positive
>103 >103 >103 >103 >103 >103 >103 >103
8 10 10 6
Positive Positive Positive Positive
>103 >103 >103 >103
Each batch contained 10 ZP-intact or 10 ZP-free embryos. Abbreviations: C-PCR, conventional polymerase chain reaction; RT-PCR, real-time polymerase chain reaction; ZP, zona pellucida.
found to be positive (4/4) and bacterial-DNA was detected in the wash baths up to the 10th wash for two batches and the eighth wash for one batch (Table 1). After 10 wash cycles, all of the exposure baths and batches of embryos were tested using RT-PCR to quantify the bacterial load. The bacterial load in the exposure baths ranged from 2.36 to 4.4 108 bacteria per mL with an average of 4.1 0.4 108 bacteria per mL. The bacterial load for embryos after the 10 wash baths was less than 104. 4. Discussion In cattle, there are no reports on the risk of C. burnetii transmission via embryo transfer. Detection of the bacterium in the genital tract of cows and bull semen is suggestive of the potential sexual transmission of Q fever, however, such transmission has yet to be confirmed [7,27]. The detection of Coxiella DNA was performed by C-PCR, which is a highly sensitive and specific method; it was shown to be capable of detecting up to 10 bacterium per mL. The quantification of bacterial load was based on RTPCR [26]. In this experiment, the probability of interaction of the bacterium with the zona pellucida and/or with the embryo cells was increased by exposing the ZP-intact and ZP-free embryos to a high concentration of Coxiella in phase I (109 bacterium per mL) over a period of 18 hours. The use of IVF embryos in this study also increased the possibility of interaction between the microorganism and the embryos in vitro, since it has been demonstrated that many pathogens adhere more readily to the surface of the zona pellucida of IVF embryos than to that of in vivo embryos [28]. Since Coxiella-DNA was detected in the first fourth wash baths for ZP-intact embryos, in all 10 wash baths for ZP-free
embryos, and also in all batches of ZP-intact and ZP-free washed embryos, this study clearly demonstrates the ability of C. burnetii to adhere to ZP-intact and ZP-free embryos. This proves that there is a risk of C. burnetii transmission after embryo transfer. Furthermore, the routine procedures proposed by the IETS are not effective for removing the bacterium from ZP-intact bovine embryos infected in vitro, although these procedures are sufficient to remove the high numbers of bacteria from the medium at the end of the exposure period. The large surface area in contact with the bacterium or the easy detachment of embryonic cells during washing may explain the presence of the bacterium up to the 10th wash bath for ZP-free embryos. Penetration of the embryonic cells is possible but multiplication is unlikely because the intracellular replication of Coxiella requires 2 days [29]. Although ZP is an effective barrier that protects embryonic cells against various pathogenic agents (viral and bacterial), our study demonstrates that C. burnetii can adhere to or penetrate the surface of the ZP and thus resist washing. This result supports our previous findings in goats demonstrating binding of Coxiella burnetii to the ZP of caprine embryos isolated from in vivo fertilized goats, after in vitro infection despite the washing procedure recommended by the IETS for bovine embryos [17]. In a previous study, Mycoplasma bovis, Mycoplasma bovigenitalium, and Ureaplasma diversum were isolated from all batches of ZPintact and ZP-free early bovine embryos after in vitro or in utero infection despite the standard IETS washing protocol, which can reduce the bacterial load associated with the ZP [21]. Our results suggest that C. burnetii is strongly bound to the ZP. The envelope of C. burnetii, similar to gram-negative bacteria, contains outer membrane proteins, A-1-gamma type peptidoglycon and lipopolysaccharide (LPS). Phase I C. burnetii (virulent form) is characterized by complete LPS, which hide the surface proteins of the external membrane [3]. LPS has an important role for entry into the host cell by engaging the integrin complex, leukocyte response integrin (avb3) and integrin-associated protein, in the infection of phase I Coxiella [3]. In contrast, the ZP is a glycoprotein membrane containing oligosaccharide chains attached to polypeptide side chains [30,31]. Although the ZP does not have an integrin complex (leukocyte response integrin and integrin-associated protein), the role of this LPS in the attachment of Coxiella to the ZP is highly plausible. Purification of the inoculum by dilution and differential centrifugation allows us to eliminate the role of egg protein [32]; the inoculum used in this study was ovoculture Coxiella. Furthermore, C. burnetii has been found in the semen of infected mice, the bacteria were mainly attached to the heads of spermatozoa via receptors on the surface of the spermatozoa [7]. These studies lead us to conclude that adhesion to the ZP is dependent on the structure of the bacterial membrane and that transmission via embryo transfer may be possible. Our results demonstrate that C. burnetii adheres to and/or penetrates the early embryonic cells and the ZP of in vitro bovine embryos after in vitro infection, and that the IETS washing protocol for bovine embryos failed to remove it. The persistence of these bacteria after washing makes the embryo a potential means of transmission of the bacterium during
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embryo transfer from infected donor cows to healthy recipients and/or their offspring. Further studies are needed (1) to determine the location of the bacterium after in vitro infection and washing by immunofluorescence, (2) to investigate whether the enzymatic and/or antibiotic treatment of infected bovine embryos would eliminate the bacterium from the ZP, and (3) to verify if similarly results are obtained with in vivo–derived embryos. Acknowledgments The authors would like to thank Véronique Blouin, Sylvie Saleun, and all technical departments at the UMR649 Institut de Recherche Thérapeutique, IRT1 INSERM, Nantes cedex 01, France. References [1] Stringfellow DA. The potential of bovine embryo transfer for infectious disease control. Rev Sci Tech Off Int Epizoot 1985;4:843–59. [2] Chalmers RM, Thomas DR, Sillis M, Softley P, Caul EO, Salmon RL, et al. Coxiella burnetii in farm workers and their families. Proc. Society for Veterinary Epidemiology and Preventive Medicine, Ed. MV Thrusfield and EA Goodall, Ennis, UK, 128–138. [3] Maurin M, Raoult D. Q fever. Clin Microbiol Rev 1999;12:518–53. [4] To H, Htwe KK, Kako N, Kim HJ, Yamaguchi T, Fukushi H, et al. Prevalence of Coxiella burnetii infection in dairy cattle with reproductive disorders. J Vet Med Sci 1998;60:859–61. [5] Bildfell RJ, Thomson GW, Haines DM, McEwen BJ, Smart N. Coxiella burnetii infection is associated with placentitis in cases of bovine abortion. J Vet Diag Invest 2000;12:419–25. [6] Berri M, Souriau A, Crosby M, Crochet D, Lechopier P, Rodolakis A. Relationships between the shedding of Coxiella burnetii, clinical signs and serological responses of 34 sheep. Vet Rec 2001;148:502–5. [7] Kruszewska D, Tylewska-Wierzbanowska S. Isolation of Coxiella burnetii bull semen. Res Vet Sci 1997;62:299–300. [8] Rousset E, Russo P, Pepin M, Raoult D. Epidémiologie de la fièvre Q animale situation en France. Med Mal Infect 2001;31:233–46. [9] Plommet M, Capponi M, Gestin J, Renoux G. Fièvre Q expérimentale des bovins. Ann Rech Vet 1973;4:325–46. [10] Guatteo R, Beaudeau F, Rodolakis A. Fièvre Q chez les bovins. Infection des bovins par Coxiella burnetii. Point Vet 2005;36:24–8. [11] Arricau-Bouvery N, Souriau A, Lechopier P, Rodolakis A. Excretion of Coxiella burnetii during an experimental infection of goats with an abortive goat strain CbC1. Ann New Ann New York Acad Sci 2003a; 990:524–6. [12] Arricau-Bouvery N, Souriau A, Lechopier P, Rodolakis A. Experimental Coxiella burnetii infection in pregnant goats: excretion routes. Vet Res 2003b;34:423–33. [13] Berri M, Crochet D, Santiago S, Rodolakis A. Spread of Coxiella burnetii in a flock of sheep after an episode of Q fever. Vet Rec 2005; 157:737–40. [14] Guatteo R, Beaudeau F, Berri M, Rodolakis A, Joly A, Seegers H. Shedding routes of Coxiella burnetii in dairy cows: implications for detection and control. Vet Res 2006;37:827–33.
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[15] Guatteo R, Beaudeau F, Joly A, Seegers H. Coxiella burnetii shedding by dairy cows. Vet Res 2007;38:849–60. [16] Alsaleh A, Pellerin JL, Rodolakis A, Larrat M, Cochonneau D, Bruyas JF, et al. Detection of Coxiella burnetii, the agent of Q fever, in oviducts and uterine flushing media and in genital tract tissues of the non pregnant goat. CIMID 2011;34:355–60. [17] Alsaleh A, Fieni F, Rodolakis A, Bruyas JF, Roux C, Larrat M, et al. Can Coxiella burnetii be transmitted by embryo transfer in goats? Theriogenology 2011;80:571–5. [18] Stringfellow DA. Recommendation for the sanitary handling of in vivo-derived embryos. In: Stringfellow DA, Seidel SM, editors. Manual of the International Embryo Transfer Society (IETS). Champaign, IL:IETS; 1998, p.79–84. [19] Bielanski AB, Surujballi O. Leptospira borgpetersenii serovar hardjo type hardjobovis in bovine embryos fertilized in vitro. Can J Vet Res 1998;62:234–6. [20] Wolfe DF, Stringfellow DA, Riddell MG, Lauerman LH, Galik PK. Adherence of Brucella ovis to preimplantation ovine ova. Theriogenology 1988;30:387–93. [21] Riddell KP, Stringfellow DA, Panagala VS. Interaction of Mycoplasma bovis and Mycoplasma bovigenitalium with preimplantation bovine embryos. Theriogenology 1989;32:633–41. [22] Bielanski A, Algire J, Randall GCB, Surujballi O. Risk of transmission of Mycobacterium avium ssp. paratuberculosis by embryo transfer of in vivo and in vitro fertilized bovine embryos. Theriogenology 2006;66:260–6. [23] Berri M, Laroucau K, Rodolakis A. 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 2000; 72:285–93. [24] Holm P, Booth P, Vajta G, Callesen H. A protein-free SOF system supplemented with amino acids, sodium citrate and myo-inositol for bovine culture. In: Report of the 13th scientific meeting of the European Embryo Transfer Association. Lyon, France. 1997 (abstract 158). [25] Lamara A, Fieni F, Mselli-Lakhal L, Chatagnon G, Bruyas JF, Tainturier D, et al. Early embryonic cells from in vivo-produced goat embryos transmit the caprine arthritis-encephalitis virus (CAEV). Theriogenology 2002b;58:1153–63. [26] Klee SR, Tyczka J, Ellerbrok H, Franz T, Linke S, Baljer G, et al. Highly sensitive real-time PCR for specific detection and quantification of Coxiella burnetii. BMC Microbiol 2006;6:1–8. [27] Rodolakis A. Chlamydiose et Fièvre Q similitudes et différences entre ces deux zoonoses. Renc Rech Ruminants 2006;13: 395–402. [28] Bielanski A. A review on disease transmission studies in relationship to production of embryos by in vitro fertilization and to related new reproductive technologies. Biotechnol Adv 1997;5:633–56. [29] Coleman SA, Fischer ER, Howe D, Mead DJ, Heinzen RA. Temporal analysis of Coxiella burnetii morphological differentiation. J Bacteriol 2004;186:7344–52. [30] Dunbar BS. Morphological, biochemical, and immunochemical characterization of the mammalian zona pellucida. In: Hartmann J, editor. Mechanism and control of fertilization. New York: Academic Press; 1983a. p. 139–75. [31] Prasad SV, Skinner SM, Carino C, Wang N, Cartwright J, Dunbar BS. Structure and function of the proteins of the mammalian zona pellucida. Cells Tissues Organs 2000;166:148–64. [32] Williams JC, Marius G, Thomas FM. Immunological and biological characterization of Coxiella burnetii, phases I and II, separated from host components. Infect Immun 1981;32:840–51.
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Risk of Coxiella burnetii transmission via embryo transfer using in vitro early bovine embryos A. Alsaleh a, F. Fieni a, *, D. Moreno a, E. Rousset b, D. Tainturier a, J.F. Bruyas a, J.L. Pellerin a a
LUNAM University, Oniris, Nantes-Atlantic National College of Veterinary Medicine, Food Science and Engineering, Sanitary Security of Reproduction Biotechnology Unit, Nantes, France b ANSES, Laboratoire de Sophia Antipolis, France
a r t i c l e i n f o
a b s t r a c t
Article history: Received 31 October 2013 Received in revised form 17 December 2013 Accepted 17 December 2013
Coxiella burnetii, an obligate intracellular bacterium of worldwide distribution, is responsible for Q fever. Domestic ruminants are the main source of infection for humans. The objectives of this study were to determine (1) whether C. burnetii would adhere to the intact zona pellucida (ZP-intact) of early in vitro–produced bovine embryos; (2) whether the bacteria would adhere to or infect the embryos (ZP-free) after in vitro infection; and (3) the efficacy of the International Embryo Transfer Society (IETS) washing protocol. One hundred and sixty, eight- to 16-cell bovine embryos produced in vitro, were randomly divided into 16 batches of 10 embryos. Twelve batches (eight ZP-intact and four ZP-free) were incubated in a medium containing C. burnetii CbB1 (Infectiologie Animale et Santé Publique, Institut National de Recherche Agronomique Tours, France). After 18 hours of incubation at 37 C and 5% CO2 in air, the embryos were washed in 10 successive baths of a PBS and 5% fetal calf serum solution in accordance with the IETS guidelines. In parallel, four batches (two ZP-intact and two ZP-free) were subjected to similar procedures but without exposure to C. burnetii to act as controls. Ten washing fluids from each batch were collected and centrifuged for 1 hour at 13,000 g. The embryos and wash pellets were tested using conventional polymerase chain reaction. C. burnetii DNA was found in all ZP-intact and ZPFree embryos after 10 successive washes. It was also detected in the first four washing fluids for ZP-intact embryos and in the 10th wash fluid for two of the four batches of ZPfree embryos. In contrast, none of the embryos or their washing fluids in the control batches were DNA positive. These results demonstrate that C. burnetii adheres to and/or penetrates the early embryonic cells and the ZP of in vitro bovine embryos after in vitro infection, and that the standard washing protocol recommended by the IETS for bovine embryos, failed to remove it. The persistence of these bacteria after washing makes the embryo a potential means of transmission of the bacterium during embryo transfer from infected donor cows to healthy recipients and/or their offspring. Further studies are required to investigate whether enzymatic and/or antibiotic treatment of bovine embryos infected by C. burnetii would eliminate the bacteria from the ZP and to verify if similarly results are obtained with in vivo–derived embryos. Ó 2014 Elsevier Inc. All rights reserved.
Keywords: Coxiella burnetii Embryo transfer International embryo transfer society Zona pellucida Bovine embryo Polymerase chain reaction
1. Introduction
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Corresponding author. Tel.: þ33 2 40687711; fax: þ33 2 40687748. E-mail address: francis.fi[email protected] (F. Fieni).
0093-691X/$ – see front matter Ó 2014 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.theriogenology.2013.12.015
Embryo transfer (ET) is used to select and diffuse genetic traits without introducing infectious disease agents. Embryos are considered to be free of specific pathogens if their donors or uterine environment are free of the agent or if
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the embryos can resist adherence and infection until the time of collection [1]. Q fever is a zoonotic endemic disease with worldwide distribution caused by an obligate intracellular bacterium, Coxiella burnetii. Domestic ruminants are the main sources for human infection [2,3]. Infection is often asymptomatic in cattle, but it may cause abortion, reproductive failure (metritis, placentitis, and infertility), and economic losses [4–6]. Infection usually occurs via inhalation of contaminated aerosols. Sexual transmission has been implicated in domestic ruminants but has yet to be proven [7,8]. The genital tract, and in particular the placenta, gravid uterus, and mammary tissue, is the preferential target for C. burnetii in pregnant animals [9,10]. Bacterial shedding after abortion has been studied in natural and experimental infections [9,11–15]. Recently, C. burnetii was detected in the genital tract flushing media (oviducts and uterine horns) and tissues of nonpregnant goats with significant bacterial loads [16]. To our knowledge, there are no reported studies on the interaction of bovine embryos and C. burnetii. A recent study conducted in our laboratory (SSBR; Oniris, Nantes, France) demonstrated that in the goat, C. burnetii has a strong tendency to cling to the zona pellucida (ZP) after in vitro infection; the washing procedure recommended by the International Embryo Transfer Society (IETS) for bovine embryos failed to remove it [17]. The IETS has established procedures to reduce the risk of pathogen transmission, which are approved for international exchanges of embryos. However, they must be tested for each pathogen and for embryos from each different species [18]. Most pathogenic agents can be eliminated by washing ZP-intact embryos. However, some pathogens can adhere to the ZP and are not removed by these procedures, resulting in contamination of the embryo and/or uterus; these include Leptospira spp., Brucella ovis, Mycoplasma bovigenitalium, and Mycobacterium avium [19–22]. Today, the detection of Coxiella-DNA in clinical samples using polymerase chain reaction (PCR) has radically changed the diagnosis of Q fever in veterinary medicine. Conventional PCR (C-PCR) and real-time PCR (RT-PCR) techniques are highly sensitive methods; they are also very specific for the detection and quantification of C. burnetii DNA in several biological samples [14,23]. To investigate the risk of C. burnetii transmission via bovine embryo transfer, our study aims to determine whether the embryonic ZP protects early embryo cells against C. burnetii infection, and whether the bacteria adhere to or infect the cells of early bovine embryos (ZPfree) after in vitro infection. We also evaluated the efficacy of the washing procedure recommended by the IETS to decontaminate bovine embryos exposed to C. burnetii in vitro. 2. Materials and methods
intraperitoneal inoculation of three OF1 mice (8 weeks) with 0.2 mL of cow placenta homogenate. The mice were killed 9 days postinoculation and their spleens were sampled and reinoculated into specific pathogen-free embryonated hen eggs. After the third passage in the chicken embryo, Coxiella burnetii was quantified using quantitative-PCR, aliquoted, and frozen at 80 C. This preparation contained 1011 bacteria per mL. To ensure purity, each aliquot used for exposures was diluted with 10 mL PBS then centrifuged twice for 15 minutes at 2000 g; the supernatant was recovered and centrifuged for 1 hour at 13,000 g. The pellet was diluted 1:100 in the exposure medium giving a final calculated concentration of 109 bacteria per mL. 2.2. In vitro production of embryo 2.2.1. In vitro maturation Oocytes were collected by aspiration via an 18-ga needle, from 3 to 8 mm diameter follicles from the ovaries of slaughtered cows. The oocytes were washed four times in HEPES-buffered Tissue Culture Medium 199 (TCM 199, M7528) and 0.4 mg/mL BSA fraction V (A9647) and then matured in 500 mL bicarbonate buffered TCM 199 (M4530) containing 60 IU/mL penicillin, 60 mg/mL streptomycin, and 10% heat-treated calf serum (CVFSVF00-01; Eurobio, Les Ulis, France) for 24 hours at 39 C in an atmosphere of 5% CO2 in humidified air. 2.2.2. In vitro fertilization After maturation, the oocytes were washed in 2 mL of HEPES-buffered TCM 199 (M7528) and 0.4 mg/mL BSA fraction V (A9647), and once in fertilization medium, IVFTyrode’s albumin lactate pyruvate medium supplemented with 6 mg/mL fatty acid-free BSA fraction V (A8806) and 1.7 IU/mL heparin (H3149) before being transferred to fertilization wells. Frozen-thawed bull spermatozoa were used to inseminate the oocytes. Motile spermatozoa were separated on a Percoll discontinuous density gradient (45%–90%) (BoviPure; Nidacon), after centrifugation at 300 g for 20 minutes at room temperature. Spermatozoa were diluted to a final concentration of 4 106 mL. Finally, oocytes and spermatozoa were incubated together for 18 hours at 39 C in a humidified atmosphere of 5% CO2 in air. 2.2.3. In vitro culture The cumulus cells were removed from the oocytes by vortexing them for 2 minutes. Presumptive zygotes were washed in PBS medium before being cultured in groups of approximately 30 embryos in 30-mL droplets of culture medium covered with mineral oil in an atmosphere of 5% O2, 5% CO2, and 90% N2 at 39 C. The culture medium was synthetic oviduct fluid, according to Holm, et al. [24], enriched with 0.4% BSA (A8806). The day of fertilization was considered as Day 0. Embryos were collected on Day-4 postinsemination for in vitro–exposure to CbB1.
2.1. Coxiella burnetii strain 2.3. Removal of the ZP The C. burnetii strain (CbB1) phase I used in this study (provided by IASP, INRA Tours, France) was isolated from the placenta of an aborted cow. It had been purified by
In accordance with the method described by Lamara, et al. [25], the embryos were placed in 1% preincubated
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pronase (Protease, P 6911; Sigma, Paris, France) at 37 C. After 60 to 90 seconds, the embryos were transferred to preincubated acidic Tyrode’s solution (pH 2.1) at 37 C for 90 to 120 seconds. Finally, the embryos were washed three times with minimum essential medium (Sigma) supplemented with 10% fetal calf serum (FCS).
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and a negative control (distilled water) were performed. Samples analyzed for C. burnetii DNA using PCR were considered positive when a band of 337 bp, corresponding to the positive control, was visualized on agarose gel electrophoresis under ultraviolet light. The sensitivity of this PCR method had been proven in our laboratory (SSBR); it detects 10 bacteria per mL of bacterial suspension (data not shown).
2.4. Experimental design 2.6. Real-time PCR procedure Four days after IVF, 160 embryos, were randomly divided into 16 batches of 10 embryos. Twelve batches (eight ZP-intact and four ZP-free) were placed in 1 mL of minimum essential medium (M2414; Sigma) supplemented with 10% FCS, 1% L-glutamine (2 mM final), 1% HEPES (0.01 M final), 2.5 mg/mL1 amphotericin B, and 50 mg/mL gentamicin and containing 109 Coxiella per mL of CbB1 strain (IASP, INRA Tours). After incubation for 18 hours at 37 C in an atmosphere of 5% CO2, the embryos were recovered and washed in batches in 10 successive baths of a PBS and 5% FCS according to the IETS guidelines [18]. A new sterile pipette was used for each successive wash; each wash corresponded to a dilution of 1:100 of the previous medium. In parallel, four batches (two ZP-intact and two ZP-free) were subjected to similar procedures but without exposure to C. burnetii as control group. All the 10 wash baths were collected separately, and centrifuged for 1 hour at 13,000 g. The washed embryos and the pellets of the 10 centrifuged wash baths were frozen at 20 C before examination for evidence of C. burnetii using PCR. 2.5. Conventional PCR procedure DNA was extracted from the batches of embryos and the wash bath pellets using a “QIAamp Blood and body fluid kit, Qiagen, France” in accordance with the manufacturer’s instructions. The detection of Coxiella-DNA was performed by amplifying a DNA fragment (337 bp) located in the transposon-like repetitive region (IS1111) gene, which is present in multiple copies in the Coxiella burnetii genome, using two primers: Trans B: 50 - CAAGAATGATCGTAACGATGCGC-30 (349–371 bp) and Trans M: 50 - CTCGTAATCACCAATCGCTTCG-30 (664–685 bp) (IASP, INRA de Tours). Three microliters of extracted DNA were added to 22 mL of amplification solution. The latter contained 5 mL of ready-to-use solution containing all reagents required for PCR: HOT FIREPol DNA polymerase, Proofreading enzyme, 5 Blend Master Mix Buffer, 7.5 mM MgCl2, 2 mM dNTPs of each, BSA, blue dye, yellow dye, and a compound to increase sample density for direct loading (Solis BioDyne, Estonia), 0.75 mL of both Trans B and Trans M primer (20 mM; Eurofins MWG Operon, Ebersberg, Germany), and 15.5 mL of distilled water DNase–RNase free. Amplification was performed in a thermal cycler (Mastercycler; Eppendorf) on the following program: after initial denaturation at 94 C for 10 minutes, the samples were subjected to a series of 35 cycles of 30 seconds denaturation at 94 C, 1 minute hybridization at 63 C, and a 3-minute elongation phase at 72 C. This was followed by a final elongation phase at 74 C for 10 minutes. Products were visualized by electrophoresis on 1.5% agarose gel. A positive control of C. burnetii (IASP, INRA Tours)
Real-time PCR was used to amplify a DNA fragment of 76 bp from the isocitrate dehydrogenase gene (icd), of which there is only one copy in the C. burnetii genome. The following primers were used: forward, icd-439F: CGT TAT TTT ACG GGT GTG CCA (439–459) and reverse, icd-514R: CAG AAT TTT CGC GGA AAA TCA (494–514), with a TaqMan probe icd-464 TM: FAM-CAT ATT CAC CTT TTC AGG CGT TTT GAC CGT-TAMRA-T (464–492) as described by Klee, et al. [26]. DNase–RNase-free water was used as a negative control. Standard series containing: 2.103, 2.104, 2.105, 2.106, 2.107 C. burnetii per mL (IASP, INRA Tours) were extracted using the QIAamp DNA mini kit (Qiagen) and used as a positive control. Five microliters of extracted DNA were added to 20 mL of RT-PCR reaction mix. The latter was composed of 12.5 mL TaqMan Universal Master Mix II (Applied Biosystems), 2.5 mL of a mixture of forward and reverse primers (0.3 mM; Eurofins MWG Operon), 0.25 mL TaqMan probe (50 nM; Eurofins MWG Operon). Water was added to make a final volume of 20 mL. All RT-PCR reactions were performed in duplicate in an ABIPRISM Sequence Detection System 7300 (Applied Biosystems) as follows: after 2 minutes at 50 C and 10 minutes at 95 C, the samples were subjected to a series of 40 cycles comprising 15 seconds at 95 C and 30 seconds at 60 C. Data were analyzed with the corresponding software. The C. burnetii titers in the samples were calculated in comparison with a standard curve obtained from a standard serial dilution of the bacteria. 2.7. Statistical analysis Fisher’s exact test was used to compare the efficacy of all the 10 embryo-washing procedures in removing Coxiella burnetii from the washing media after in vitro infection. Values of P less than 0.05 were considered to be significant. 3. Results In total, 160 ZP-intact and ZP-free embryos were assigned to 18 batches. Twelve batches were exposed in vitro to Coxiella CbB1 at a concentration of 109 Coxiella per mL; the remaining six batches were used as controls. Coxiella-DNA was detected by C-PCR in all batches of infected ZP-intact and ZP-free embryos (8/8 and 4/4 respectively) after being washed 10 times (Table 1). Bacterial DNA was not detected in any of the embryos in the control batches. For ZP-intact embryo the first three washing media of the infected group were consistently found to be positive (8/8), whereas subsequent washes tested negative (P < 0.05). For ZP-free embryo the first six washing media of the infected group were consistently
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Table 1 Detection of Coxiella burnetii in successive embryo wash baths and batches of infected ZP-intact eight- to 16-cell embryos and infected ZP-free eightto 16-cell embryos after 10 wash cycles, using C-PCR and quantification of Coxiella burnetii in embryo exposure baths and in batches of infected ZPintact eight- to 16-cell embryos or infected ZP-free eight- to 16-cell embryos by RT-PCR. Batch of Exposure embryos bath (Coxiella per mL) (RT-PCR)
Zona pellucida intact 1 2.6 108 2 2.5 108 3 2.4 108 4 2.8 108 5 2.9 108 6 4.4 108 7 4.4 108 8 4.4 108 Zona pellucida free 1 2.9 108 2 2.9 108 3 2.9 108 4 2.9 108
Last positive wash for C. burnetii (C-PCR)
Batch of embryos after 10 wash baths Detection of C burnetii (C-PCR)
Quantification of C burnetii (RT-PCR)
4 3 4 3 3 4 3 5
Positive Positive Positive Positive Positive Positive Positive Positive
>103 >103 >103 >103 >103 >103 >103 >103
8 10 10 6
Positive Positive Positive Positive
>103 >103 >103 >103
Each batch contained 10 ZP-intact or 10 ZP-free embryos. Abbreviations: C-PCR, conventional polymerase chain reaction; RT-PCR, real-time polymerase chain reaction; ZP, zona pellucida.
found to be positive (4/4) and bacterial-DNA was detected in the wash baths up to the 10th wash for two batches and the eighth wash for one batch (Table 1). After 10 wash cycles, all of the exposure baths and batches of embryos were tested using RT-PCR to quantify the bacterial load. The bacterial load in the exposure baths ranged from 2.36 to 4.4 108 bacteria per mL with an average of 4.1 0.4 108 bacteria per mL. The bacterial load for embryos after the 10 wash baths was less than 104. 4. Discussion In cattle, there are no reports on the risk of C. burnetii transmission via embryo transfer. Detection of the bacterium in the genital tract of cows and bull semen is suggestive of the potential sexual transmission of Q fever, however, such transmission has yet to be confirmed [7,27]. The detection of Coxiella DNA was performed by C-PCR, which is a highly sensitive and specific method; it was shown to be capable of detecting up to 10 bacterium per mL. The quantification of bacterial load was based on RTPCR [26]. In this experiment, the probability of interaction of the bacterium with the zona pellucida and/or with the embryo cells was increased by exposing the ZP-intact and ZP-free embryos to a high concentration of Coxiella in phase I (109 bacterium per mL) over a period of 18 hours. The use of IVF embryos in this study also increased the possibility of interaction between the microorganism and the embryos in vitro, since it has been demonstrated that many pathogens adhere more readily to the surface of the zona pellucida of IVF embryos than to that of in vivo embryos [28]. Since Coxiella-DNA was detected in the first fourth wash baths for ZP-intact embryos, in all 10 wash baths for ZP-free
embryos, and also in all batches of ZP-intact and ZP-free washed embryos, this study clearly demonstrates the ability of C. burnetii to adhere to ZP-intact and ZP-free embryos. This proves that there is a risk of C. burnetii transmission after embryo transfer. Furthermore, the routine procedures proposed by the IETS are not effective for removing the bacterium from ZP-intact bovine embryos infected in vitro, although these procedures are sufficient to remove the high numbers of bacteria from the medium at the end of the exposure period. The large surface area in contact with the bacterium or the easy detachment of embryonic cells during washing may explain the presence of the bacterium up to the 10th wash bath for ZP-free embryos. Penetration of the embryonic cells is possible but multiplication is unlikely because the intracellular replication of Coxiella requires 2 days [29]. Although ZP is an effective barrier that protects embryonic cells against various pathogenic agents (viral and bacterial), our study demonstrates that C. burnetii can adhere to or penetrate the surface of the ZP and thus resist washing. This result supports our previous findings in goats demonstrating binding of Coxiella burnetii to the ZP of caprine embryos isolated from in vivo fertilized goats, after in vitro infection despite the washing procedure recommended by the IETS for bovine embryos [17]. In a previous study, Mycoplasma bovis, Mycoplasma bovigenitalium, and Ureaplasma diversum were isolated from all batches of ZPintact and ZP-free early bovine embryos after in vitro or in utero infection despite the standard IETS washing protocol, which can reduce the bacterial load associated with the ZP [21]. Our results suggest that C. burnetii is strongly bound to the ZP. The envelope of C. burnetii, similar to gram-negative bacteria, contains outer membrane proteins, A-1-gamma type peptidoglycon and lipopolysaccharide (LPS). Phase I C. burnetii (virulent form) is characterized by complete LPS, which hide the surface proteins of the external membrane [3]. LPS has an important role for entry into the host cell by engaging the integrin complex, leukocyte response integrin (avb3) and integrin-associated protein, in the infection of phase I Coxiella [3]. In contrast, the ZP is a glycoprotein membrane containing oligosaccharide chains attached to polypeptide side chains [30,31]. Although the ZP does not have an integrin complex (leukocyte response integrin and integrin-associated protein), the role of this LPS in the attachment of Coxiella to the ZP is highly plausible. Purification of the inoculum by dilution and differential centrifugation allows us to eliminate the role of egg protein [32]; the inoculum used in this study was ovoculture Coxiella. Furthermore, C. burnetii has been found in the semen of infected mice, the bacteria were mainly attached to the heads of spermatozoa via receptors on the surface of the spermatozoa [7]. These studies lead us to conclude that adhesion to the ZP is dependent on the structure of the bacterial membrane and that transmission via embryo transfer may be possible. Our results demonstrate that C. burnetii adheres to and/or penetrates the early embryonic cells and the ZP of in vitro bovine embryos after in vitro infection, and that the IETS washing protocol for bovine embryos failed to remove it. The persistence of these bacteria after washing makes the embryo a potential means of transmission of the bacterium during
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embryo transfer from infected donor cows to healthy recipients and/or their offspring. Further studies are needed (1) to determine the location of the bacterium after in vitro infection and washing by immunofluorescence, (2) to investigate whether the enzymatic and/or antibiotic treatment of infected bovine embryos would eliminate the bacterium from the ZP, and (3) to verify if similarly results are obtained with in vivo–derived embryos. Acknowledgments The authors would like to thank Véronique Blouin, Sylvie Saleun, and all technical departments at the UMR649 Institut de Recherche Thérapeutique, IRT1 INSERM, Nantes cedex 01, France. References [1] Stringfellow DA. The potential of bovine embryo transfer for infectious disease control. Rev Sci Tech Off Int Epizoot 1985;4:843–59. [2] Chalmers RM, Thomas DR, Sillis M, Softley P, Caul EO, Salmon RL, et al. Coxiella burnetii in farm workers and their families. Proc. Society for Veterinary Epidemiology and Preventive Medicine, Ed. MV Thrusfield and EA Goodall, Ennis, UK, 128–138. [3] Maurin M, Raoult D. Q fever. Clin Microbiol Rev 1999;12:518–53. [4] To H, Htwe KK, Kako N, Kim HJ, Yamaguchi T, Fukushi H, et al. Prevalence of Coxiella burnetii infection in dairy cattle with reproductive disorders. J Vet Med Sci 1998;60:859–61. [5] Bildfell RJ, Thomson GW, Haines DM, McEwen BJ, Smart N. Coxiella burnetii infection is associated with placentitis in cases of bovine abortion. J Vet Diag Invest 2000;12:419–25. [6] Berri M, Souriau A, Crosby M, Crochet D, Lechopier P, Rodolakis A. Relationships between the shedding of Coxiella burnetii, clinical signs and serological responses of 34 sheep. Vet Rec 2001;148:502–5. [7] Kruszewska D, Tylewska-Wierzbanowska S. Isolation of Coxiella burnetii bull semen. Res Vet Sci 1997;62:299–300. [8] Rousset E, Russo P, Pepin M, Raoult D. Epidémiologie de la fièvre Q animale situation en France. Med Mal Infect 2001;31:233–46. [9] Plommet M, Capponi M, Gestin J, Renoux G. Fièvre Q expérimentale des bovins. Ann Rech Vet 1973;4:325–46. [10] Guatteo R, Beaudeau F, Rodolakis A. Fièvre Q chez les bovins. Infection des bovins par Coxiella burnetii. Point Vet 2005;36:24–8. [11] Arricau-Bouvery N, Souriau A, Lechopier P, Rodolakis A. Excretion of Coxiella burnetii during an experimental infection of goats with an abortive goat strain CbC1. Ann New Ann New York Acad Sci 2003a; 990:524–6. [12] Arricau-Bouvery N, Souriau A, Lechopier P, Rodolakis A. Experimental Coxiella burnetii infection in pregnant goats: excretion routes. Vet Res 2003b;34:423–33. [13] Berri M, Crochet D, Santiago S, Rodolakis A. Spread of Coxiella burnetii in a flock of sheep after an episode of Q fever. Vet Rec 2005; 157:737–40. [14] Guatteo R, Beaudeau F, Berri M, Rodolakis A, Joly A, Seegers H. Shedding routes of Coxiella burnetii in dairy cows: implications for detection and control. Vet Res 2006;37:827–33.
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