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REVIEW
10.1111/1469-0691.12163
Relevance of Rift Valley fever to public health in the European Union
V. Chevalier
UPR Animal et Gestion Integree des Risques (AGIRs), CIRAD, Montpellier, France
Abstract
Rift Valley fever (RVF), a vector-borne zoonotic disease caused by a phlebovirus (family Bunyaviridae), is considered to be one of the most
important viral zoonoses in Africa. It is also a potential bioterrorism agent. Transmitted by mosquitoes or by direct contact with viraemic
products, RVF affects both livestock and humans, causing abortion storms in pregnant ruminants and sudden death in newborns. The disease
provokes flu syndrome in most human cases, but also severe encephalitic or haemorrhagic forms and death. There is neither a treatment
nor a vaccine for humans. The disease, historically confined to the African continent, recently spread to the Arabian Peninsula and Indian
Ocean. Animal movements, legal or illegal, strongly contribute to viral spread, threatening the Mediterranean basin and Europe, where
competent vectors are present. Given the unpredictability of virus introduction and uncertainties about RVF epidemiology, there is an
urgent need to fill the scientific gaps by developing large regional research programmes, to build predictive models, and to implement early
warning systems and surveillance designs adapted to northern African and European countries.
Keywords: Europe, public health, Rift Valley fever, risk, surveillance, trade
Clin Microbiol Infect
Corresponding author: V. Chevalier, Campus International de
Baillarguet, 34398, Montpellier, France
E-mail: [email protected]
Rift Valley fever (RVF) is a vector-borne zoonotic disease
caused by a phlebovirus (family Bunyaviridae). RVF virus (RVFV)
is an enveloped RNA virus characterized by a genome
composed of three segments, designated L, M, and S, of
negative or ambisense polarity [1]. Like many bunyaviruses,
RVFV produces a non-structural protein encoded by the S
segment, the NSs protein, which acts as a virulence factor [2].
It is transmitted from ruminants to ruminants by mosquito
bites, mainly from the genera Aedes and Culex, but also from
the genera Anopheles and Mansonia, as recently suggested in
Madagascar and Kenya [3,4]. Direct transmission between
ruminants through contact with viraemic fluids, i.e. blood or
fetal liquid, is also strongly suspected. Humans are mostly
contaminated after contact with aborted fetal material, i.e.
placental membranes from infected ruminants, which contain
large numbers of virus particles and blood. Furthermore, RVFV
was observed or experimentally demonstrated to persist for
long periods in different biotic or abiotic settings: a laboratory
assistant was infected in a laboratory 4 months after the virus
was handled in this laboratory; the virus may be isolated from
carcase tissues such as spleen or liver between 36 and 72 h
after death; and infected sheep plasma retained RVFV
infectivity after 8 years of storage and shipment under a
variety of refrigeration conditions [5–8]. Consequently, veterinarians and laboratory, agricultural and slaughterhouse workers may be at risk. If it exists, the viral load in raw milk is
assumed to be low. The presence of virus in nasal and
lachrymal secretions and the urine and faeces of infected
animals has not been demonstrated [1,9]. To date, no humanto-human transmission of RVF has been documented.
The health and economic consequences of RVF outbreaks
are severe. Besides losses resulting from animal trade control,
RVFV infection causes abortion storms in pregnant ruminants
and acute deaths in newborns. However, the severity of clinical
signs depends on the species: sheep are more susceptible than
goats, which are themselves more susceptible than cattle and
camels. In adults, one may observe non-specific signs such as
vomiting, diarrhoea, respiratory disease, fever, lethargy, and
anorexia [10]. Although, in the majority of human cases, RVFV
causes a mild illness with fever, headache, myalgia, and liver
abnormalities, a minority of human cases of infection may lead
to either retinitis with permanent vision loss, encephalitis, or
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Clinical Microbiology and Infection ª2013 European Society of Clinical Microbiology and Infectious Diseases
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Clinical Microbiology and Infection
haemorrhagic forms that may lead to death [11]. During the
2000 Saudi Arabia outbreak, the major clinical characteristics
reported among 165 consecutive patients included a high
frequency (75%) of hepatocellular failure, acute renal failure for
42% of patients, and haemorrhagic manifestations for approximately 19% of patients. A total of 56 patients died (33.9%)
[12]. There is no aetiological treatment, for either animals or
humans. Several vaccines are under development, but, to date,
there are no licensed and commercially available vaccines to
protect humans [13]. Regarding ruminants, the ‘Smithburn’
vaccine, a live attenuated vaccine, has been used for years in
Africa. It cheaply and efficiently protects sheep and cattle with
a single inoculation, but it may cause abortion or teratogenic
effects in fetuses, and may present a risk of reversion to
virulence: its use is thus reduced to endemic areas. A new,
promising live attenuated vaccine candidate—clone 13—was
obtained from a strain isolated from a mild human case in the
Central African Republic [14,15]. This vaccine was recently
registered and marketed in South Africa [16]. Owing to its
severity, RVFV is considered to be a major zoonotic threat to
the USA, and is number 3 on the list of the 17 most dangerous
animal threats, behind highly pathogenic avian influenza and
food and mouth disease [17].
Early detection and implementation of appropriate measures, which are essential to minimize the consequences of
outbreaks, require a deep understanding of transmission,
spread and persistence mechanisms. However, the epidemiology of RVF is complex. The disease is enzootic in many African
countries and Madagascar, with outbreaks occurring every 5–
15 years. However, the factors triggering outbreaks and the
way in which the virus persists during inter-epizootic periods
remain mostly unknown. RVF has been reported in four
epidemiological systems:
1. ‘Dambo’ areas, in East Africa. Dambos are shallow depressions that can be 1 km in length and several hundreds of
metres in width, and are often located in valleys near rivers.
In these areas, a correlation between heavy rainfall events
and RVF outbreak occurrence has been clearly demonstrated. Viral transmission from one Aedes mcintoshi mosquito generation to another by ‘vertical transmission’, and
the survival of infected eggs in dry mud for several years,
could explain the maintenance of the virus in the field during
inter-epizootic periods [18,19].
2. Semi-arid areas of western Africa—Senegal and Mauritania
—characterized by temporary areas of water. In these
areas, a recent modelling study showed that outbreaks
could not be directly related to heavy rainfall events, but
mostly to abundant regular rainfall occurring throughout the
rainy season, which is favourable for successive high density
of the main vectors, i.e. Aedes vexans and Culex poicilipes [20].
The persistence of the virus may result from either the
above-mentioned vertical transmission in A. vexans mosquitoes or from the regular introduction of the virus by
nomadic herds [21].
3. Irrigated areas such as the Nile Delta or Senegal river basin,
where permanent water may favour Culex population
persistence, and thus RVFV transmission throughout the
year [22–24].
4. Temperate and mountainous areas, as recently demonstrated in Madagascar, where transmission and spread result
from local vector-borne transmission associated with
specific cattle trade habits [25,26].
A role of wild ruminants, which is strongly suspected in
southern Africa, needs further investigation [27].
RVFV was historically confined to the African continent until
2000, when it was reported for the first time in the Arabian
Peninsula [28]; the geographical distribution of the virus has
recently increased. Global changes, including climatic changes,
may be involved. However, the natural history of RVF shows
that animal movements, legal or illegal, have strongly contributed to the spread of the virus [29–31]. Infected mosquitoes,
travelling in aircraft or cargoe, could also be incriminated [32].
Unprecedented increases in the international trade and
worldwide movements of humans, animals and animal products
are thus likely to alter the epidemiological patterns of RVF. In
fact, there is a large livestock trade between the sub-Saharan
countries where the virus is circulating and northern African
countries. The 2010 and 2012 Mauritanian outbreaks [33,34],
associated with the recent detection of serologically positive
camels in Morocco coming from the southern part of the
Saharan desert in a north-western direction [35], demonstrated RVFV in northern Africa. Because of the illegal
importation of ruminants and the short geographical distance
between southern European coasts and northern Africa, the
exposure of the Mediterranean basin and Europe to RVFV has
increased. According to the European Food Safety Authority,
the virus could be introduced by either legally or illegally
imported infected animals, infected vectors, legally or illegally
imported contaminated animal products, fomites, or vaccines;
the first of these is the most plausible [36]. Even if this
probability is considered to be very low, because trading in
livestock from northern Africa and the Middle East to Europe is
forbidden, this introduction into an area of a dense and na€ıve
ruminant population may be devastating [37]. Conversely, and
because the virus can circulate with few or even no clinical
signs, it could remain undetected and settle in endemic foci in
areas where eco-climatic conditions are favourable. In fact, 50
mosquito species may transmit the virus, and some of them are
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Clinical Microbiology and Infection ª2013 European Society of Clinical Microbiology and Infectious Diseases, CMI
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Chevalier
present in Europe [36,37]. Among them, Culex pipiens, whose
European distribution is wide, is competent to transmit RVFV
[38]. The distribution of Aedes albopictus, which is another
potential vector of RVFV, has dramatically enlarged since its
first introduction [39,40]. Established homogeneous populations have been identified in Albania, Croatia, France, Greece,
Monaco, Montenegro, Italy, San Marino, Slovenia, and Spain
[41]. The changing European climate could facilitate this spread
to new areas [42], enlarging the distribution of areas suitable
for RVFV transmission.
RVF should be suspected when a sudden abortion storm or
sudden deaths of ruminants are associated or not with febrile
syndrome in humans. Depending on the epidemiological status
of the area, and the delay post-infection, diagnosis may be
performed either by detection of live virus, viral antigen or viral
nucleic acids within 1–10 days after the onset of the disease, or
by detection of acute-phase (IgM) or chronic (IgG) antibodies,
starting from 4 days post-infection [1]. Among recently validated tests, a sandwich ELISA for antigen detection (sAg-ELISA)
was recently reported [43], having, respectively, 67.7% and 70%
sensitivity for humans and sheep, and 97.9% and 100% specificity,
and real-time reverse transcriptase isothermal amplification
assays (RT-LAMP) have been developed and tested, allowing the
detection of a wide spectrum of isolates and in clinical specimens
in 30 min [44]. Inhibition ELISA tests for detecting IgG in all
species, capture ELISA for IgM for bovines, caprines and ovines
and sandwich ELISA for IgG for the same species are commercially available. Finally, the virus neutralization test, which is
considered to be the reference standard, is highly accurate, with
no or few cross-reactions with other phleboviruses [45,46].
However, this methodology requires live virus, and thus can be
used only in biosafety level 3 laboratories [1].
Several control options are available, such as vaccines in
animals, larvicides in vector breeding sites and/or insecticide
spraying, animal trade control, and the provision of information
to exposed human populations. However, the disease is usually
well established in animal populations by the time when the first
human cases are observed [13]: in endemic areas, animal
vaccination is probably the best way to protect human health.
Regarding surveillance and virus-free areas, the use of a
dense sentinel herd network for surveillance would be costprohibitive unless strictly focused on ecologically defined risky
areas. Syndromic surveillance relies on the early detection of
abnormal clusters of illness indicators rather than clinical signs,
and thus reduces the time-lag between the onset of the
outbreak and the diagnosis [47,48]. This methodology may be
a useful alternative in the case of RVF, which may provoke
non-specific signs, in either animals or humans: RVF human
cases were detected in 2009 in Mayotte thanks to the
surveillance for dengue-like syndromes [49].
Relevance of Rift Valley fever to public health
3
In the Horn of Africa, RVF outbreaks can successfully be
predicted with lead times of 2–4 months, thanks to remotely
sensed data-driven models [50]. This early warning system is
based on accurate knowledge of the disease epidemiology. As
far as Europe is concerned, there is an urgent need to fill
scientific gaps, i.e. to experimentally evaluate European
potential vector competence and European ruminant breed
susceptibility, and to assess the existence of ruminant to
ruminant direct transmission. For implementation of a riskbased surveillance network, European areas that are potentially suitable for virus transmission need to be identified [51]:
given the current lack of knowledge, the multi-criteria decision
analysis method could be a valuable tool allowing the
integration of expert knowledge, the available literature and
data with trade—legal or illegal—information [52]. Furthermore, the epidemiological situation in northern African
countries, and the risk of introduction via either animal
movements or infected vector ‘travel’, should be assessed, as
well as the performance of both existing northern African and
European surveillance systems. In fact, a‘one-health’ regional
approach and a joint effort by human and animal health
authorities is needed to control RVF in endemic countries and
protect virus-free areas from introduction of the virus.
Transparency Declaration
The author declares having no conflict of interest related to
the present article.
References
1. Pepin M, Bouloy M, Bird BH, Kemp A, Paweska JT. Rift Valley fever
(Bunyaviridae: Phlebovirus): an update on pathogenesis, molecular
epidemiology, vectors, diagnostics and prevention. Vet Res 2010; 41: 61.
2. Billecocq A, Spiegel M, Vialat P, et al. NSs protein of Rift Valley fever
virus blocks interferon production by inhibiting host gene transcription.
J Virol 2004; 78: 9798–9806.
3. Ratovonjato J, Olive M, Tantely L et al. Detection, isolation, and genetic
characterization of Rift Valley fever virus from Anopheles (Anopheles)
coustani, Anopheles (Anopheles) squamosus, and Culex (Culex) antennatus of the Haute Matsiatra Region, Madagascar. Vector Borne Zoonotic
Dis 2010; 11: 753–759.
4. Sang RC, Ahmed O, Faye O et al. Entomologic investigations of a
chikungunya virus epidemic in the Union of the Comoros. Am J Trop
Med Hyg 2008; 78: 77–82.
5. Andrewes C, Horstman D. The susceptibility of viruses to ethyl ether.
J Gen Microbiol 1949; 3: 290–297.
6. Craig D, Thomas W, DeSanctis A. Stability of Rift Valley fever virus at
4 °C. Appl Microbiol 1967; 15: 446–447.
7. Francis T, McGill T. Rift Valley fever. A report of 3 cases of laboratory
infection and the experimental transmission of the disease to ferrets.
J Exp Med 1935; 62: 433–438.
8. Easterday B. Rift valley fever. Adv Vet Sci 1965; 10: 65–127.
ª2013 The Author
Clinical Microbiology and Infection ª2013 European Society of Clinical Microbiology and Infectious Diseases, CMI
4
CMI
Clinical Microbiology and Infection
9. Easterday BC, Murphy LC, Benett DG. Experimental Rift Valley fever in
lambs and sheep. Am J Vet Res 1962; 23: 1231–1240.
10. Gerdes G. Rift Valley fever. Rev Sci Tech Off Int Epiz 2004; 23: 613–623.
11. LaBeaud A, Muiruri S, Sutherland LJ et al. Postepidemic analysis of Rift
Valley fever virus transmission in northeastern Kenya: a village cohort
study. PLoS Negl Trop Dis 2011; 5: e1265.
12. Al-Hazmi M, Ayoola EA, Abdurahman M et al. Epidemic Rift Valley
fever in Saudi Arabia: a clinical study of severe illness in humans. Clin
Infect Dis 2003; 36: 245–252.
13. Breiman R, Minjauw B, Sharif S, Ithondeka P, Njenga MK. Rift Valley
fever: scientific pathways toward public health prevention and
response. Am J Trop Med Hyg 2010; 83(suppl 2): 1–4.
14. Dungu B, Louw I, Lubisi A, Hunter P, von Teichman B, Bouloy M.
Evaluation of the efficacy and safety of the Rift Valley Fever Clone 13
vaccine in sheep. Vaccine 2010; 28: 4581–4587.
15. Muller R, Saluzzo JF, Lopez N et al. Characterization of clone 13, a
naturally attenuated avirulent isolate of Rift Valley fever virus, which is
altered in the small segment. Am J Trop Med Hyg 1995; 53: 405–411.
16. Kortekaas J, Zingeser J, De Leeuw P, Rocque DL, Unger H, Moormann
R. Rift Valley fever vaccine development, progress and constraints.
Emerg Infect Dis 2011; 17.
17. Mandell R, Flick R. Rift Valley fever virus: a real bioterror threat.
J Bioterror Biodef 2011; 2: 108.
18. Linthicum KJ, Davies FG, Kairo A, Bailey CL. Rift Valley fever virus
(family Bunyaviridae, genus Phlebovirus). Isolations from Diptera collected
during an inter-epizootic period in Kenya. J Hyg 1985; 95: 197–209.
19. Linthicum K, Anyamba A, Tucker C, Kelley P, Myers M, Peters C.
Climate and satellite indicators to forecast Rift Valley fever epidemics
in Kenya. Science 1999; 285: 397–400.
20. Soti V, Tran A, Degenne P et al. Combining hydrology and mosquito
population models to identify the drivers of Rift Valley fever
emergence in semi-arid regions of West Africa. PLoS Negl Trop Dis
2012; 6: e1795.
21. Chevalier V, Lancelot R, Thiongane Y, Sall B, Diaite A, Mondet B. Rift
Valley fever in small ruminants, Senegal, 2003. Emerg Infect Dis 2005; 11:
1693–1700.
22. Meegan J. The Rift Valley fever epizootic in Egypt 1977–78. 1.
Description of the epizootic and virological studies. Trans Roy Soc Trop
Med Hyg 1979; 73: 618–623.
23. El-Akkad AM. Rift Valley fever outbreak in Egypt. October–December
1977. J Egypt Public Health Assoc 1978; 53: 123–128.
24. Digoutte JP, Peters CJ. General aspects of the 1987 Rift Valley fever
epidemic in Mauritania. Res Virol 1989; 140: 27–30.
25. Chevalier V, Rakotondrafara T, Jourdan M et al. An unexpected
recurrent transmission of Rift Valley fever virus in cattle in a temperate
and mountainous area of Madagascar. PLoS Neg Trop Dis 2011; 5: 1–9.
26. Nicolas G, Durand B, Duboz R, Rakotondravao R, Chevalier V.
Description and analyses of trading network of cattle on Madagascar
highlands and potential role in the diffusion of Rift Valley fever virus.
Acta Trop 2012; 126: 19–27. Available at: http://dx.doi.org/10.1016/j.
actatropica.2012.12.013.
27. Olive M, Goodman S, Reynes J. The role of wild mammals in the
maintenance of Rift Valley fever virus. J Wildl Dis 2012; 48: 241–266.
28. Ahmad K. More deaths from Rift Valley fever in Saudi Arabia and
Yemen. Lancet 2000; 356: 1422.
29. Abd El-Rahim IHA, El-Hakim UA, Hussein M. An epizootic of Rift Valley
fever in Egypt in 1997. Rev Sci Tech Off Int Epi 1999; 18: 741–748.
30. Shoemaker T, Boulianne C, Vincent MJ et al. Genetic analysis of viruses
associated with emergence of Rift Valley fever in Saudi Arabia and
Yemen, 2000–2001. Emerg Infect Dis 2002; 8: 1415–1420.
31. Carroll S, Reynes J, Khristova M, Andriamandimby S. Genetic evidence
for Rift valley fever outbreaks in Madagascar resulting from virus
introductions from the East African mainland rather than enzootic
maintenance. J Virol 2011; 95: 6162–6167.
32. Russell R. Survival of insects in the wheel bays of a Boeing 747B aircraft
on flights between tropical and temperate airports. Bull World Health
Organ 1987; 65: 659–662.
33. El Mamy A, Baba M, Barry Y et al. Unexpected Rift Valley fever
outbreak, northern Mauritania. Emerg Infect Dis 2011; 17: 1894–1896.
34. World Health Organization. Rift Valley fever in Mauritania, D.O.N.
WHO Global Alert and Response (GAR), Editor 2012, WHO.
Available at: http://www.who.int/csr/don/2012_11_01/en/index.html
35. El-Harrak M, Martin-Folgar R, Llorente F et al. Rift Valley and West
Nile virus antibodies in camels, North Africa. Emerg Infect Dis 2011; 17:
2372–2374.
36. European Food Safety Authority. The risk of Rift Valley fever incursion and
its persistence within the Community, in EFSA journal 2005. Parma: EFSA,
2005; 1–128.
37. Chevalier V, Pepin M, Plee L, Lancelot R. Rift Valley fever—a threat for
Europe? Euro Surveill 2010; 15: 1–11.
38. Moutailler S, Krida G, Schaffner F, Vazeille M, Failloux AB. Potential
vectors of Rift Valley fever virus in the Mediterranean region. Vect Born
Zoo Dis 2008; 8: 749–753.
39. Moutailler S, Bouloy M. Short report: effcient oral infection of Culex
pipien quinquefasciatus by RVF virus using a cotton stick support. Am J
Trop Med Hyg 2007; 76: 827–829.
40. Turell MJ, Bailey CL, Beaman JR. Vector competence of a Houston,
Texas strain of Aedes albopictus for Rift Valley fever virus. J Am Mosq
Control Assoc 1988; 4: 94–96.
41. European Centre for Disease Prevention and Control. Areas of
possible establishment of Aedes albopictus (the tiger mosquito) in
Europe for 2010 and 2030. In: Impacts of Europe’s changing climate—
2008 indicator-based assessment. Stockolm: ECDC, 2008.
42. Roiz D, Neteler M, Castellani C, Arnoldi D, Rizzoli A. Climatic factors
driving invasion of the tiger mosquito (Aedes albopictus) into new areas
of Trentino, northern Italy. PLoS ONE 2011; 6: 1–8.
43. Jansen van Vuren P, Potgieter A, Paweska J, van Dijk A. Preparation and
evaluation of a recombinant Rift Valley fever virus N protein for the
detection of IgG and IgM antibodies in humans and animals by indirect
ELISA. J Virol Methods 2007; 140: 106–114.
44. Le Roux C, Kubo T, Grobbelaar A et al. Development and evaluation
of a real-time reverse transcription-loop-mediated isothermal amplification assay for rapid detection of Rift Valley fever virus in clinical
specimens. J Clin Microbiol 2009; 47: 647–651.
45. Swanepoel R, Struthers JK, Erasmus MJ et al. Comparison of
techniques for demonstrating antibodies to Rift Valley fever virus.
J Hyg (Lond) 1986; 97: 317–329.
46. Tesh RB, Peters CJ, Meegan JM. Studies on the antigenic relationship
among phleboviruses. Am J Trop Med Hyg 1982; 31: 149–155.
47. Chu A, Savage R, Willison D et al. . The use of syndromic surveillance
for decision-making during the H1N1 pandemic: a qualitative study.
BMC Public Health 2012; 12: 929.
48. Leblond A, Hendrikx P, Sabatier P. West Nile virus outbreak detection
using syndromic monitoring in horses. Vector Borne Zoonotic Dis 2007;
7: 404–410.
49. InVS. Bulletin de veille sanitaire N°02/Novembre 2009. InVS: La Reunion,
France, 2009.
50. Anyamba A, Linthicum K, Small J et al. Prediction, assessment of the
Rift Valley fever activity in East and Southern Africa 2006–2008
and possible vector control strategies. Am J Trop Med Hyg 2010; 83
(suppl 2): 43–51.
51. St€ark KD, Regula G, Hernandez J et al. Concepts for risk-based
surveillance in the field of veterinary medicine and veterinary public
health: review of current approaches. BMC Health Serv Res 2006; 28: 6–
20.
52. Boroushaki S, Malczewski J. Using the fuzzy majority approach for GISbased multicriteria group decision-making. Comput Geosci 2010; 36:
302–312.
ª2013 The Author
Clinical Microbiology and Infection ª2013 European Society of Clinical Microbiology and Infectious Diseases, CMI