CSIRO PUBLISHING

Reproduction, Fertility and Development, 2015, 27, 63–71 http://dx.doi.org/10.1071/RD14367

Viral emergence and consequences for reproductive performance in ruminants: two recent examples (bluetongue and Schmallenberg viruses) Ste´phan Zientara A,C and Claire Ponsart B A

UPE, ANSES, INRA, ENVA, UMR 1161 ANSES/INRA/ENVA, Laboratoire de sante´ animale d’Alfort, 23 Avenue du Ge´ne´ral de gaulle, 94703 Maisons-Alfort, France. B ANSES, Unite´ des zoonoses bacte´riennes, Laboratoire de sante´ animale d’Alfort, 23 Avenue du Ge´ne´ral de gaulle, 94703 Maisons-Alfort, France. C Corresponding author. Email: [email protected]

Abstract. Viruses can emerge unexpectedly in different regions of the world and may have negative effects on reproductive performance. This paper describes the consequences for reproductive performance that have been reported after the introduction to Europe of two emerging viruses, namely the bluetongue (BTV) and Schmallenberg (SBV) viruses. Following the extensive spread of BTV in northern Europe, large numbers of pregnant cows were infected with BTV serotype 8 (BTV-8) during the breeding season of 2007. Initial reports of some cases of abortion and hydranencephaly in cattle in late 2007 were followed by quite exhaustive investigations in the field that showed that 10%–35% of healthy calves were infected with BTV-8 before birth. Transplacental transmission and fetal abnormalities in cattle and sheep had been previously observed only with strains of the virus that were propagated in embryonated eggs and/or cell culture, such as vaccine strains or vaccine candidate strains. After the unexpected emergence of BTV-8 in northern Europe in 2006, another arbovirus, namely SBV, emerged in Europe in 2011, causing a new economically important disease in ruminants. This new virus, belonging to the Orthobunyavirus genus in the Bunyaviridae family, was first detected in Germany, in The Netherlands and in Belgium in 2011 and soon after in the UK, France, Italy, Luxembourg, Spain, Denmark and Switzerland. Adult animals show no or only mild clinical symptoms, whereas infection during a critical period of gestation can lead to abortion, stillbirth or the birth of severely malformed offspring. The impact of the disease is usually greater in sheep than in cattle. The consequences of SBV infection in domestic ruminants and more precisely the secondary effects on off-springs will be described. Additional keywords: abortion, infertility.

Introduction Many different agents, both infectious and non-infectious, are capable of crossing the placenta to cause fetal injury. Most maternal virus infections are not transmitted to the fetus; however, several viruses that infect domestic animals have the capacity to cross the placenta and subsequently induce disease and/or developmental defects (teratogenesis). The outcome of viral infections of the fetus depends on the susceptibility of the fetus to the infecting virus, which, in turn, is dependent on the gestational age of the fetus at the time of exposure as well as the virulence of the virus. The potential consequences of fetal viral infections include teratogenesis, fetal disease with or without abortion, growth retardation, persistent postnatal infection or no obvious abnormality (MacLachlan et al. 2000). The effects of emerging viruses on reproductive performance are described for two viruses that have emerged in Europe Journal compilation Ó IETS 2015

recently, namely bluetongue virus (BTV) serotype 8 (BTV-8) and Schmallenberg virus (SBV). Development of bluetongue in Europe Bluetongue (BT) is a non-contagious, insect-transmitted disease of certain breeds of sheep and some species of wild ruminants (e.g. red deer) that is caused by BTV (Verwoerd and Erasmus 2004). BTV infection of ruminants occurs throughout much of the temperate and tropical regions of the world, coincident with the distribution of specific species of Culicoides biting midges that are biological vectors for the virus (Gibbs and Greiner 1994; Tabachnick 2004). BT typically occurs when susceptible sheep are introduced into areas where virulent strains of BTV circulate, or when virulent strains of BTV extend their range into previously unexposed populations of ruminants. The global www.publish.csiro.au/journals/rfd

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distribution of BTV has historically been between latitudes of approximately 40–508N and 358S. However, during the recent northern European epidemic, the virus spread far beyond its prior known upper northern limits (Savini et al. 2008). BTV is the prototype member of the genus Orbivirus, family Reoviridae (Mertens et al. 2004). All reoviruses share distinct common properties, including segmented genomes of doublestranded (ds) RNA and characteristic virion morphology and structure. To date, 26 distinct BTV serotypes have been described that all share common group antigens but are distinguished on the basis of serotype-specific virus neutralisation assays. Importantly, there is considerable variation among field strains of BTV, even those of the same serotype, which reflects differences in the nucleotide sequence of each of the 10 distinct dsRNA segments of the BTV genome (Bonneau et al. 1999; Pritchard et al. 2004). Genetic heterogeneity of field strains of BTV occurs as a result of both genetic drift and genetic shift, the latter resulting from the reassortment of viral genes during mixed infections of either the vertebrate or invertebrate hosts of the virus (Bonneau et al. 2001; Bonneau and MacLachlan 2004). However, unlike most single stranded (ss) RNA viruses, the orbiviruses are genetically and antigenically stable throughout infection; point mutations do not appear to arise in vivo, at least at the high frequency noted with many non-segmented ssRNA viruses (Schwartz-Cornil et al. 2008). BTV protection is serotype specific. Immunisation with one of the 26 BTV serotypes does not elicit a high cross-protection against other serotypes. After inoculation by a biting midge and initial replication in the lymph nodes draining the sites of inoculation, BTV disseminates to secondary sites, principally the lungs and spleen, where it replicates in the endothelium and mononuclear phagocytes. BTV can disseminate via the lymph and/or blood. BTV infects monocytes both in vivo and in vitro. In vivo, infectious BTV can be retrieved transiently (,1 week) from monocytes (minimum 105 cells; Barratt-Boyes and 1994). Monocytes also express BTV antigens in vivo at low frequency (four non-structural (NS2) antigen-positive monocytes per 2
105 peripheral blood mononuclear cells). Conversely, resting T lymphocytes are not efficient at supporting BTV replication unless they are activated by mitogens. Interestingly, gdT cell lines can be efficiently infected in vitro and blood gdT cells from infected sheep (3–13 days after infection) have been induced to produce infectious BTV when cocultured with skin fibroblasts (Schwartz-Cornil et al. 2008). However, it is unclear how monocytes and possibly blast T cells are involved in vivo in the pathogenesis of BTV. Last but not least, infectious BTV can also be detected in the intracellular vesicles of erythrocytes, where it does not replicate but persists in invaginations of the cell membrane (Schwartz-Cornil et al. 2008). The association of infectious BTV with erythrocytes is detected very early after infection (24 h) and persists throughout viraemia. Consequently, BTV infection in ruminants is characterised by a prolonged cell-associated viraemia that can persist in the presence of high titres of neutralising antibody, although recovered animals are immune to re-infection with the homologous serotype of BTV. In sheep and cattle, infectious BTV can be detected in the blood for 35–60 days and the viral genome can be detected up to 160 days (Schwartz-Cornil et al. 2008).

S. Zientara and C. Ponsart

Five different BTV serotypes (1, 2, 4, 9 and 16) have recently spread throughout extensive areas of Mediterranean Europe, and BTV-8 emerged in northern Europe in 2006 (Toussaint et al. 2006). Before 2006, BT epidemics had been reported, although irregularly, from 1979 to 1999 in Greece. In 1999, BT was reported in mainland Greece and south-eastern Bulgaria (Calistri et al. 2004). BTV-2 emerged in 2000 in France (Corsica), Italy (Sardinia) and Spain (Balearic Islands; Zientara et al. 2002; Savini et al. 2008) and BTV-9 appeared later during the same year in Italy. In 2002, BTV-4 and BTV-16 were detected in the southern regions of Italy, and in 2003 BTV-4 emerged in Sardinia, Corsica and Menorca; in 2003, BTV-16 was reported in Sardinia and then, in 2004, in Corsica (Bre´ard et al. 2004). In 2006, BTV-8 emerged very unexpectedly in the north of Europe, affecting Belgium, France, Germany, Luxembourg and The Netherlands (Savini et al. 2008). Subsequent to the spread of BTV-1, -2, -4, -9 and -16 within the Mediterranean basin, BTV-6, -8 and -11 all appeared in northern Europe (Benelux) after 2006. Although BTV-6 and -11 are closely related to South African live attenuated vaccine viruses, the origin of the highly pathogenic strain of BTV-8 that has now spread throughout much of Europe remains unknown (MacLachlan and Guthrie 2010). Since its appearance in northern Europe, this virus spread to the Mediterranean basin, Scandinavia and the Middle East. This strain of BTV-8 exhibits several distinctive features that are unusual among other field strains of BTV, notably its ability to cross the ruminant placenta with high frequency. More importantly, certain live attenuated BTV vaccine strains, passaged extensively in cell culture, were observed to be capable of transplacental transmission in both cattle and sheep (MacLachlan and Guthrie 2010), but early studies with wildtype (WT) BTV did not reveal this property (Acree et al. 1991; Roeder et al. 1991). However, unexpectedly, following the introduction of the pathogenic BTV-8 into northern Europe in 2006, it was observed that transplacental transmission of this virus strain occurred in both cattle and sheep (Rasmussen et al. 2013). The particular features of the European BTV-8 virus that are responsible for transplacental transmission are not known. However, this property of the virus and the subsequent birth of viraemic offspring may contribute its ability to be maintained from one year to the next (overwintering) within northern Europe in the absence of an active vector population during the winter months. Vaccination is central to prevention of BT in many endemic areas, as well as to the response to incursions of the disease into previously unaffected regions. There is little cross-protection between BTV serotypes, so to achieve comprehensive protection animals should be vaccinated against all BTV serotypes that circulate in a given region. Inactivated vaccines have been used since 2005 in some European countries (including France and Italy; Schwartz-Cornil et al. 2008). However, the available inactivated vaccines are directed against only a few serotypes. Effects of BTV infection on gametes and embryos Very little virus is found in the secretions and excretions of infected animals and the disease is not normally regarded as

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3 inseminations 12 h apart, recovery on Day 6 following AI 1 donor - 7 embryos; 1 donor - 0 embryo Detection of virus in BT cell cultures (cytopathic effect/ electron microscopy): positive in blood, cervix, uterine horn, ovary of both donor cows No virus on or in embryos –

Ewes remained uninfected through 42 days after exposure (no seroconversion) 11

11

Hare et al. (1988)

Schlafer et al. (1990)

Positive bull

2 seronegative superovulated cows

12 uninfected anoestrous ewes

11 Hare et al. (1988)

2 BTV-infected rams

9 days before breeding, 2
105 CEIVD50 of a BTV suspension (from BTV-infected midges) (TCID50 105.8/0.1 mL)

35 Day 4 embryos (4 unfertilised eggs), 19 seronegative cross-bred ewes – 31 anoestrous ewes (inoculated 2 days before breeding) 9 days before breeding, 105 CEIVD50 of a BTV suspension (from BTV-infected midges)

17 Thomas et al. (1985)

Positive

?

9 heifers inseminated with semen from a shedding bull 4 seronegative superovulated heifers ? 20 bulls 10, 11, 13, 17

BTV-infected rams

20 Day 6 transferred embryos, 16 seronegative recipients

Virus isolated from 19 of 181 ejaculates (seminal shedding) Virus never isolated from most (13/20) bulls 6 of 9 heifers became viraemic and seroconverted 2 donor heifers infected by the semen (viraemia) - 9 of 10 pregnancies, 1 abortion: 6 of 7 live calves, 1 of 2 dystocia No seroconversion observed in recipients or calves No virus isolated from flushing media Pregnancies in 11 recipients No seroconversion observed in recipients or lambs (n ¼ 12)

Results Embryo transfer Donors Male inoculation Male status Serotype Reference

contagious by the oral route, or via aerosols (European Food Safety Authority 2011). However, BTV can be demonstrated in the semen of viraemic bulls (Howard et al. 1985; Mu¨ller et al. 2010). In case of natural infection, it has been reported that semen quality (assessed on the basis of motility and the percentage of abnormalities) decreased in rams (n ¼ 78; Kirschvink et al. 2008) and bulls (Foster et al. 1980; Mu¨ller et al. 2010). Field results from 78 rams contaminated with BTV-8 showed considerable variability between rams and reported that normal semen quality may be restored approximately 85 days after the clinical phase (Kirschvink et al. 2008). It must be noted that viruses are still infectious after freezing and then thawing. The vaccines against BTV-1 and -8 used in Europe are inactivated and have no effect on semen quality (Savini et al. 2008). With regard to in vivo-produced embryos, the use of semen infected with different BTV serotypes will often lead to contamination of donor cows (Thomas et al. 1985; Schlafer et al. 1990; Table 1). Following insemination with infected semen, the virus has been isolated from the blood, cervix, uterine horn and ovaries of donor cows (Schlafer et al. 1990). A few experiments in cattle (Thomas et al. 1985; Schlafer et al. 1990) and small ruminants (Hare et al. 1998) have evaluated the possibility that BTV may be transmitted via infected semen to in vivo-produced embryos. These studies have shown that, at least in the case of embryos washed according to the International Embryo Transfer Society (IETS) protocols, the virus is not transmitted to recipients or newborns, despite the use of contaminated semen and the induction of viral infection in inseminated cows or contamination of the embryos (Table 1). Results in sheep (Gilbert et al. 1987; Singh et al. 1997), goats (Ali Al Ahmad et al. 2012) and cows (Singh et al. 1982; Schlafer et al. 1990; De Clercq et al. 2008) confirmed that embryos exposed in vitro to the virus can induce an intrauterine infection leading to viraemia and seroconversion of the recipients after embryo transfer. However, many experiments have been conducted with naturally infected donor goats and cows (Acree et al. 1991), in addition to experimentally inoculated ewes (Hare et al. 1988; Singh et al. 1997) and cows (Thomas et al. 1983; Acree et al. 1991). In these experiments, no virus was isolated either from the flushing fluids (except for one ewe; Singh et al. 1997) or from the embryos. No seroconversion of recipients or newborns was observed. These studies evaluated BTV-4, -10, -11, -17 and -18 in ewes and cows, and BTV-6 and -14 in goats. On the basis of these experiments, BTV has been listed in IETS Category 1 (i.e. it is a ‘disease agent for which there is sufficient evidence to show that the risk of transmission is negligible provided that the embryos are properly handled between collection and transfer’; see Appendix 3.3.5 of the OIE Terrestrial Animal Health Code, http://www.oie.int/fr/ normes-internationales/code-terrestre/). Other studies have investigated the effects of BTV on in vitro-produced embryos (Ali Al Ahmad et al. 2011, 2012; Vandaele et al. 2012). BTV appears to have a very high affinity for the zona pellucida of in vitro-exposed embryos, adhering in large numbers to its surface (Ali Al Ahmad et al. 2011, 2012) and being able to replicate in blastocyst cells

Reproduction, Fertility and Development

Table 1. Contamination of embryos arising from fertilisation with virus infected semen BTV, bluetongue (BT) virus; TCID50, 50% tissue culture infective dose; CEIVD50, 50% chicken embryo intravascular lethal dose of BTV

Viral emergence and reproduction in ruminants

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(Vandaele et al. 2012). However, no embryo transfer experiments have been performed. Effects of BTV infection on reproductive performance (abortions) The incursion of BTV-8 in northern France in 2007 gave rise to clinical signs never seen before in sheep and cattle on the French mainland (nasal discharge, swelling of the head and neck, conjunctivitis, swelling inside and ulceration of the mouth, swollen teats, tiredness, saliva drooling out of the mouth; Le Gal et al. 2008). An increased number of abortions in cattle was observed in the affected regions, but the cause was not clearly established. Initial reports of some cases of abortion and hydranencephaly in cattle in late 2007 were followed by quite exhaustive investigations in the field that showed that 10%–35% of healthy calves were infected with BTV-8 before birth (European Food Safety Authority 2011). More precisely, during the winter of 2007–08, an outbreak of unprecedented developmental lesions of the central nervous system (CNS) was detected in newborn or stillborn calves and lambs among the routine submissions to the Faculty of Veterinary Medicine, Lie`ge, Belgium, for postmortem examination (Desmecht et al. 2008). The congenital malformations ranged from brains with thinwalled cerebral hemispheres, variably sized cerebral cysts and dilated lateral ventricles (porencephaly with some degree of hydrocephalus) to brains in which the cerebral hemispheres were represented only by fluid-filled sacs (hydranencephaly). The brain stem structures were usually present, but the cerebellum was sometimes dysplasic, cystic or replaced by a fluid-filled sac. The observation of a series of hydranencephalopathies in calves and lambs examined postmortem (and the simultaneous increase in anti-BTV-8 antibody titres measured in paired sera from a cohort of precolostral calves and their dams) illustrated that the BTV-8 strain that spread across Europe in 2007 differed from previous WT strains in that it was able to cross the ruminant placenta spontaneously and infect the fetus (Desmecht et al. 2008). Studies conducted in Belgium and The Netherlands indicated that BTV-8 transplacental infection could occur in cattle (De Clercq et al. 2008; Desmecht et al. 2008; Wouda et al. 2008; Backx et al. 2009). Recent peer-reviewed publications from several European research groups have reported transplacental transmission of BTV-8 in sheep and cattle in the field (European Food Safety Authority 2011). Publications describing animal trials with BTV-8 are limited. In these studies, the proportion of transplacentally infected offspring of the total number of offspring tested from infected dams varies considerably (e.g. from 0% to 37% seropositive offspring born from infected dams). Previous peer-reviewed publications from Australian and American researchers (European Food Safety Authority 2011) have also examined transplacental transmission of BTV-1, -4, -10, -11, -13, -16, -17, -20, -21, -22, -23 and undetermined serotypes. In two of these reports (Kirkbride and Johnson 1989), no transplacental transmission of undefined field BTV was reported in a large number of cattle (n ¼ 994) and sheep (n ¼ 553). In another

S. Zientara and C. Ponsart

10 studies describing animal trials with experimental infections with BTV strains other than BTV-8, the BTV used mostly had a passage history in eggs and/or cell lines (European Food Safety Authority 2011). Transplacental transmission of laboratory strains of BTV was observed (European Food Safety Authority 2011). Paradoxically, two studies reported transplacental transmission of field strains of BTV-11 (Kirkbride and Johnson 1989); this remains unexplained (European Food Safety Authority 2011). Giovanni Savini (Italy) examined fetal spleens and/or brains of 663 sheep, 429 cattle, 155 goats and 17 buffaloes from 804 herds in central–southern Italy from fetuses of dams vaccinated with BTV-2 or BTV-9 modified live virus (MLV) vaccines. The BTV vaccine strains were isolated from 31 fetuses (G. Savini, pers. comm.; MacLachlan et al. 2009). James MacLachlan reported that MLV vaccines are used in sheep in the US and in South Africa, and viruses genetically similar or identical to these vaccine viruses circulate in the field. A low incidence of fetal abnormalities in cattle, consistent with in utero infection with BTV, is observed and these fetuses are seropositive for BTV; however, it has not been proved whether vaccine viruses that naturally circulate are responsible for these rare fetal infections. In summary, the observed incidence of transplacental transmission was zero or very low in areas affected by BTV before the appearance of BTV-8. The low level of transplacental transmission of strains other than BTV-8 in the field seems to be related to vaccination with live-attenuated vaccines (MacLachlan and Guthrie 2010). The genetic determinants responsible for transplacental transmission of BTV have not been identified (i.e. which of the 10 gene segments or combinations thereof are involved) for either BTV-8, MLV vaccine strains or laboratory-adapted strains. Because strains of BTV readily exchange (reassort) gene segments during mixed infections of either animals or insects with more than one virus strain and/or serotype (European Food Safety Authority 2011), there is a possibility that reassortment in the field will lead to the emergence of novel virus strains or serotypes other than BTV-8 with new features (e.g. the ability to cross the placenta and/or to cause disease in cattle). Concerning pathogenesis, BTV infection of bovine and ovine microvascular endothelial cells induces the transcription of interleukin (IL)-1, IL-8, IL-6, cyclo-oxygenase-2 and inducible nitric oxide synthase (de Maula et al. 2002). These mediators are involved in the pathogenesis of severe viral haemorrhagic fevers (de Maula et al. 2002). Infection of sheep and cattle with BTV induces increases in plasma concentrations of prostacyclin and thromboxane (de Maula et al. 2002). Thromboxane is a strong procoagulant factor, whereas prostacyclin is a potent vasodilator and inhibitor of platelet aggregation. There is a much higher prostacyclin : thromboxane ratio in cattle than in sheep, which may explain the lower sensitivity of cattle to BTV-induced microvascular injury and thrombosis. The lesions in BTV-infected animals reflect virus-mediated injury to small blood vessels. BTV replicates in endothelial cells, causing cell injury and necrosis and leading to vascular thrombosis, tissue infarction and haemorrhage (SchwartzCornil et al. 2008).

Viral emergence and reproduction in ruminants

Emergence of SBV in Europe In 2011, another virus emerged in Europe. SBV was first detected in Germany and in The Netherlands in 2011. Infected adult ruminants may show no or only unspecific mild clinical signs, such as fever, diarrhoea or reduced milk production for a few days (Hoffmann et al. 2012). In December 2011, The Netherlands reported a teratogenic effect of SBV in sheep with the birth of malformed lambs with a crooked neck, hydrocephalus and stiff joints (Doceul et al. 2013). The presence of SBV was then reported in Belgium at the end of December 2011 and in the UK on 22 January 2012. France reported its first case of SBV on 25 January 2012 after the virus genome was detected by reverse transcription–polymerase chain reaction (RT-PCR) in brain samples from malformed lambs born (Doceul et al. 2013). By the end of April 2012, SBV had been detected in 3628 herds in Europe (Doceul et al. 2013). SBV-infected holdings recorded up to this date corresponded to infections occurring in 2011. In May 2012, acute SBV infections were detected in cattle in south-west France, indicating that SBV was able to recirculate after the winter period (Sailleau et al. 2013). Similar conclusions were also reached in the UK after the detection of the virus in newborn lambs born in May and June 2012 and in Germany (Doceul et al. 2013). By the end of October 2012, SBV infection was confirmed by RT-PCR and/or serology in approximately 6000 holdings in Europe (Doceul et al. 2013). These reports indicate that the virus has spread rapidly across vast parts of Europe. It must be noted that SBV has never been reported elsewhere in the world. Much more important than its effect on adult ruminants is the induction by SBV infection of severe congenital malformations, premature birth or stillbirth or the birth of mummified fetuses when the dam is infected during a critical phase of gestation. Analysis of viral sequences has led to classification of SBV in the Bunyaviridae family and the Orthobunyavirus genus. The Bunyaviridae family is comprised of 350 viruses divided into five genera: Orthobunyavirus, Hantavirus, Nairovirus, Phlebovirus and Tospovirus. Viruses such as Rift Valley fever virus (Phlebovirus), Akabane virus (Orthobunyavirus) and Nairobi sheep disease virus (Nairovirus) are important pathogens in veterinary medicine. Other viruses, such as Hantaviruses responsible for haemorrhagic fever with renal syndrome and cardiopulmonary syndrome or Crimean–Congo haemorrhagic fever virus (Nairovirus), can cause serious disease in humans (Doceul et al. 2013). The Orthobunyavirus genus is comprised of more than 170 viruses, including SBV. The bunyavirus genome consists of three segments of negative-sense ssRNA, namely the L (large), M (medium) and S (small) segments. The L segment encodes the RNA-dependent RNA polymerase, the M segment encodes the precursor of the GN and GC envelope glycoproteins and the non-structural protein NSm and the S segment encodes the nucleoprotein N and the non-structural protein NSs in an overlapping open reading frame (Hoffmann et al. 2012). The pathogenicity of orthobunyaviruses is dependent on multiple viral factors encoded by these three genomic segments. Initial phylogenetic analysis of SBV genomic segments indicated that SBV exhibits 69% identity with Akabane virus for the L segment, 71% identity with Aino virus for the M

Reproduction, Fertility and Development

67

segment and 97% identity with Shamonda virus for the S segment (Hoffmann et al. 2012). After analysis of additional sequence data, it was reported that the M segments of the Sathuperi and Douglas orthobunyaviruses exhibit higher identity with SBV, whereas the S and L segments were closer to Shamonda virus (Hoffmann et al. 2012). All these viruses belong to the Simbu serogroup, although no cross-protection has been reported between them. Most bunyaviruses are transmitted by arthropod vectors, in particular mosquitoes, phlebotoms, culicoides, ticks and thrips, with the exception of hantaviruses, which are transmitted by rodents. Studies have shown that viruses within the Simbu serogroup are mostly transmitted by culicoides, as well as by mosquitoes from the Aedes and Culex genera and by several species of ticks (Saeed et al. 2001). Recently, a study reported the presence of the SBV genome in a pool of culicoides (Culicoı¨des obsoletus and Culicoı¨des dewulfi) trapped in October 2011 in Belgium (Doceul et al. 2013). These studies suggest that these midges can act as vectors for the transmission of SBV, as they do for BTV. Sheep and goats seem to be very mildly affected by SBV infection. Symptoms are more apparent in adult cows, and include loss of appetite, hyperthermia and diarrhoea, which can lead to a 50% reduction in milk production (Gibbens 2012). Symptoms usually disappear within a few days. The viraemia induced by SBV is short lived, lasting for 2–6 days in cattle (Hoffmann et al. 2012). SBV is quite similar to another orthobunyavirus (i.e. Akabane virus) that is known to have negative effects on reproductive performance in cattle. Akabane virus is a culicoides-borne orthobunyavirus that is teratogenic to the fetus of cattle and small ruminant species (Kirkland et al. 1988). Depending on the stage of gestation at which infection occurs, and the length of gestation of the mammalian host, a range of congenital defects may be observed. The developing CNS is usually the most severely affected, with hydranencephaly and arthrogyposis most frequently observed. Less commonly, some strains of Akabane virus can cause encephalitis in the neonate or, rarely, adult cattle (Kirkland et al. 1988). Akabane viruses are known to be widespread in temperate and tropical regions of Australia, south-east Asia, the Middle East and some African countries. Disease is infrequently observed in regions where this virus is endemic and the presence of the virus remains unrecognised in the absence of serological surveillance. In some Asian countries, vaccines are used to minimise the occurrence of disease (Kirkland et al. 1988). Effects of SBV infection on gametes and embryos From initial field data, SBV-RNA was detected in only 55 semen batches of 1719 samples tested in seropositive bulls (3%; ProMed-mail 2012a, 2012b). Following experimental infection, the highest SBV-RNA concentrations in semen were observed between 4 and 7 days after infection (Van Der Poel et al. 2014). In that study, viable SBV was only isolated from blood samples and not from semen or genital tissues. Large variability has been reported in the excretion of SBV in semen of naturally infected bulls (Ponsart et al. 2014). Outcomes of virus detection in semen are influenced by the extraction method used

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S. Zientara and C. Ponsart

Table 2. Reproductive disorders reported in female ruminants infected with Schmallenberg virus SBV, Schmallenberg virus; RT-PCR, reverse transcription–polymerase chain reaction; VNT, virus neutralisation test; d.p.i., days post infection; d.g. days of gestation Reproductive disorder

Country

Species

Reference

15 lambs and 2 calves with malformations

Germany

Cattle

Bilk et al. (2012)

Netherlands

Cattle, sheep, goats

Loeffen et al. (2012)

RT-PCR test on brains of .600 lambs and 1200 calves with arthrogryposis syndrome: SBV confirmed in 22% of lambs and 19% of calves

Netherlands

Cattle, sheep

van Maanen et al. (2012)

2166 malformed newborns from 1621 farms

Netherlands

Cattle, sheep, goats

Bouwstra et al. (2013)

Belgium

Cattle, sheep

De Regge et al. (2013)

Belgium

Sheep

Saegerman et al. (2013)

Netherlands

Sheep

Stockhofe et al. (2013)

Germany

Cattle

Wernike et al. (2014)

Presence of viral genome in the placental fluid, brain, umbilicus and spinal cord of most of the animals Infected herds: 95% (190/201) of ewes and 99% (145/146) of cows seropositive Herds with malformations: .90% (231/255) of ewes and 95% (795/834) of cows seropositive

Positive RT-PCR results in 29% of lamb brains tested, 14% of calf brains tested, and 9% of goat kid brains tested RT-PCR of various tissues (brain, cerebellum, brain stem, spinal cord, thymus, spleen, lymph nodes, meconium) from 90 malformed lambs and 81 malformed calves The brain stem is the most suitable tissue to detect SBV using VNT: 95% of lambs and 44% of calves positive 26 ovine suckler herds: 13 infected (I; 961 sheep) vs 13 negative (N; 331 sheep) Increased abortion rate in I vs N herds (6.7% vs 3.2%), more malformed lambs born at term in I vs N herds (6.7% vs 3.2%) and more dystocia in I vs N herds (6.7% vs 3.2%) 21 SBV seronegative ewes inoculated with 1 mL SBV viraemic calf serum at 38 or 45 d.g. Viraemia detected 3 d.p.i. in all ewes, except one Necropsy samples taken from 39 fetuses at 7 d.p.i. Positive umbilical cord and fetal amnion fluid samples in fetuses infected at both 38 d.g. (85% and 15%, respectively) and 45 d.g. (74% and 11%, respectively) 27 calves whose mothers were naturally infected during the first 5 months of pregnancy (between 47 and 162 d.g.): 1 calf with typical malformation, 1 stillborn, 9 clinically healthy calves with high SBV antibody titres measured before colostrum intake (6 with viral RNA in meconium swabs)

(Hoffmann et al. 2013). It has been demonstrated that Trizol is highly efficient for the extraction of RNA from matrices with a potentially high amount of PCR inhibitors (Vanbinst et al. 2010; Hoffmann et al. 2013). Four different patterns of viral excretion in semen have been characterised: (1) sustained and prolonged SBV genome in consecutive semen batches, up to 2.5 months following seroconversion in rare cases (Hoffmann et al. 2013; Ponsart et al. 2014); (2) a single positive semen batch (Hoffmann et al. 2013; Steinrigl et al. 2013); (3) intermittent excretion patterns (Hoffmann et al. 2013; Van Der Poel et al. 2014); or (4) the absence of SBV-RNA in the semen (Hoffmann et al. 2013; Ponsart et al. 2014). Effects of SBV infection on reproductive performance (abortions and teratogenic effects) In December 2011, The Netherlands reported a teratogenic effect of SBV in sheep with manifestations comparable to those

observed for Akabane and Aino viruses (Elbers et al. 2012). Infected females were able to transmit the virus to fetuses (ovine, caprine and bovine), which developed atypical malformations leading most frequently to intrauterine death or death immediately after birth (Table 2). Common congenital malformations and clinical signs in aborted and stillborn animals include a neuromusculoskeletal disorder called arthrogryposis, severe torticollis, ankylosis, kyphosis, lordosis, scoliosis, brachygnathia inferior and neurological disorders such as amaurosis, ataxia and/or behavioural abnormalities (‘dummy syndrome’, as observed during the epizooty caused by BTV-8; Doceul et al. 2013). Newborns suffer from severe neurological disorders that generally lead to the death of the animal between several hours and several days after birth. It was reported that a 1-week-old SBV-positive calf born at term had severe CNS lesions, severe dysfunction of the cerebral cortex, basal ganglia and mesencephalon, severe porencephaly or hydranencephaly (lack of

Viral emergence and reproduction in ruminants

brain cerebral hemispheres) but no arthrogryposis (van den Brom et al. 2012). Interestingly, the SBV genome was still detectable in the CNS, suggesting that it is able to persist in the infected fetus after birth (Table 2). Necropsy has revealed some cases of hydranencephaly, hydrocephaly, cerebral and cerebellar hypoplasia and porencephaly in sheep and cattle offspring (Garigliany et al. 2012; Hahn et al. 2013; van den Brom et al. 2012). In the case of Akabane virus, infection of the fetus occurs between Days 28 and 36 of gestation in the ovine, Days 30 and 50 in the caprine and Days 76 and 174 in the bovine (Kirkland et al. 1988). The severity of fetal injury depends on the time of infection during gestation, with maximal damage for infections that occur when neuronal tissues are differentiating (Kirkland et al. 1988). In all probability, this critical period is similar for SBV. However, not every infection of the dam with SBV or other Simbu serogroup viruses during that time frame results in an abnormal course of gestation or leads to a fetal antibody response (Table 2; Wernike et al. 2014). Conclusion The ability of the emerged European strain of BTV-8 to cross the ruminant placenta has been established in experimental and field studies in both sheep and cattle. In the first year after the recognised introduction of SBV into north-west Europe, musculoskeletal malformations and pathological changes of the CNS, such as porencephaly, hydranencephaly and hypoplasia of the cerebellum, in newborn lambs and calves were the most intriguing clinical features of this infection. Therefore, SBV joins the group of other teratogenic, arthroborne viruses, such as Akabane virus. On the basis of epidemiological studies of recent SBV outbreaks and comparisons with the pathogenesis of Akabane virus, it is assumed that the teratogenic infection takes place during the first trimester; however, the efficiency of transplacental infection generally and in relation to the gestation time point is unknown. In addition, information on the transplacental transfer and viral tropism in the uterus and fetus is lacking. Many questions remain regarding the pathogenesis of these viral infections in pregnant animals, their vertical transmission (by embryos and/or gametes), their effects on fertility, the dynamics of the virus towards and in the fetus. In light of such unexpected emergence, it is necessary that specialists in the field of reproduction and virologists collaborate quickly, as in the case of the BTV-8 and SBV infections. References Acree, J. A., Echternkamp, S. E., Kappes, S. M., Luedke, A. J., Holbrook, F. R., Pearson, J. E., and Ross, G. S. (1991). Failure of embryos from bluetongue infected cattle to transmit virus to susceptible recipients of their offspring. Theriogenology 36, 689–697. doi:10.1016/0093-691X (91)90406-4 Ali Al Ahmad, M. Z., Pellerin, J. L., Larrat, M., Chatagnon, G., Ce´cile, R., and Sailleau, C. (2011). Can bluetongue virus (BTV) be transmitted via caprine embryo transfer? Theriogenology 76, 126–132. doi:10.1016/ J.THERIOGENOLOGY.2011.01.025 Ali Al Ahmad, M. Z., Bruyas, J. F., Pellerin, J. L., Larrat, M., Chatagnon, G., Roux, C., Sailleau, C., Zientara, S., and Fieni, F. (2012). Evaluation

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