Transboundary and Emerging Diseases
ORIGINAL ARTICLE
Molecular and Serological Studies on the Rift Valley Fever Outbreak in Mauritania in 2010 €ckel1, M. Eiden1, B. O. EL Mamy2, K. Isselmou2, A. Vina-Rodriguez1, B. Doumbia3 and S. Ja M. H. Groschup1 1 2 3
Institute for Novel and Emerging Infectious Diseases, Friedrich-Loeffler-Institut, Greifswald - Insel Riems, Germany Service de Pathologie Infectieuses, Centre National de l’Elevage et de Recherches Veterinaires (CNERV), Nouakchott, Mauritanie Minist ere du d eveloppement rural, Nouakchott, Mauritania
Keywords: Rift Valley fever virus; small ruminants; camels; Mauritania Correspondence: M. H. Groschup. Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of Novel and Emerging Infectious €dufer 10, 17493 Greifswald - Isle Diseases, Su of Riems, Germany. Tel.: +49 38351 71163; Fax: +49 38351 71191; E-mail: martin. [email protected] Received for publication November 15, 2012 doi:10.1111/tbed.12142
Summary Rift Valley fever virus (RVFV) is a vector-borne RNA virus affecting humans, livestock and wildlife. In October/November 2010, after a period of unusually heavy rainfall, a Rift Valley fever outbreak occurred in northern Mauritania causing clinical cases in cattle, sheep, goats and camels, 21 of which were of lethal outcome. The aim of this study was to obtain further information on the continuation of RVF virus activity and spread in animal species in Mauritania after this outbreak. We therefore tested sera from small ruminants, cattle and camels for the presence of viral RNA and antibodies against RVFV. These sera were collected in different parts of the country from December 2010 to February 2011 and tested with three different ELISAs and an indirect immunofluorescence assay. The results show a high seroprevalence of RVFV IgM and IgG antibodies of about 57% in all animals investigated. Moreover, in four camel sera, viral RNA was detected emphasizing the important role camels played during the latest RVF outbreak in Mauritania. The study demonstrates the continuous spread of RVFV in Mauritania after initial emergence and highlights the potential role of small ruminants and camels in virus dissemination.
Introduction Rift Valley fever virus (RVFV) is a zoonotic arbovirus affecting humans and a wide range of vertebrate hosts such as small ruminants, cattle, camels and wild ruminants. Rift Valley fever virus was first discovered in 1931 in Kenya (Great Rift Valley) in a huge outbreak, which claimed the lives of more than 4500 lambs and ewes (Daubney et al., 1931; referenced in Bird et al., 2009). Rift Valley fever virus is a single-stranded RNA virus belonging to the family Bunyaviridae; genus Phlebovirus. Its genome consists of three segments, the M (medium)- and L (large)-segment are of negative orientation whereas the S (small)-segment has ambisense polarity. The S-segment encodes the nucleocapsid protein (N-protein) and a non-structural protein Nss. The M-segment encodes the two glycoproteins Gn and Gc and also includes the genetic information for a nonstructural protein Nsm, whereas the L-segment codes for the RNA-dependent RNA-polymerase (Pepin et al., 2010).
Numerous mosquito species are considered to be competent RVFV transmission vectors, several of them can also be found in Europe (Moutailler et al., 2008; Pepin et al., 2010). Rift Valley fever virus infections in humans usually lead to a self-limiting febrile illness. However, in 1–2% of these infections, humans develop retinitis, encephalitis, haemorrhagic fever and acute hepatitis, sometimes with lethal outcome. Morbidity and mortality in livestock are much higher, as infections are characterized by high mortality rates of up to 100% among newborn lambs and by high abortion rates, especially among pregnant sheep and goats, the so-called ‘abortion storms’. Rift Valley fever virus transmission to humans can occur by direct contact with infected body fluids, tissues and/or excretions or via mosquito bites (Bird et al., 2009; Bouloy and Weber, 2010). The infection risk is especially high after periods of long and heavy rainfalls, as mosquitoes in suddenly flooded puddles and river beds hatch from infected eggs in which
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the virus can survive over many years. Blood-feeding female mosquitoes transmit RVFV to animals, which act as first amplification host and eventually infect humans (Bird et al., 2009). Until 1977, RVF outbreaks were restricted to sub-Saharan Africa, but more recently cases have also been found in Egypt (Ahmed Kamal, 2011) and Sudan (Gad et al., 1986). The first outbreak outside of the African continent was in 2000/2001 in Saudi Arabia and Yemen. Several outbreaks have been reported in different African countries since 2006: Sudan in 2007 (Seufi and Galal, 2007;; Hassan et al., 2011), Kenya in 2006/2007 (Munyua et al., 2010; Nguku et al., 2010; Hightower et al., 2012), Tanzania in 2007 (Swai and Schoonman, 2009; Jost et al., 2010) and Madagascar in 2008/9 (Chevalier et al., 2011; Jeanmaire et al., 2011). In 2010, more outbreaks were found in South Africa, Namibia, Saudi Arabia, Botswana and Mauritania (OIE; WAHID interface) with a total of more than 14 500 clinical cases and over 9000 dead livestock and wild animals. In Mauritania and Namibia, outbreaks resumed in early 2011, whereas cases in South Africa continued throughout the year. The first RVF cases were seen in Mauritania in 1987 when 220 humans died from the disease (Saluzzo et al., 1987, 1989; Jouan et al., 1988; Digoutte and Peters, 1989; Ksiazek et al., 1989). More RVF epidemics occurred subsequently in 1993 (Zeller et al., 1995), 1998 (Nabeth et al., 2001), 2003 (Faye et al., 2007), 2010 (El Mamy et al., 2011) and 2012 (Promedmail, 2012). Outbreaks in 2010 were concentrated on three regions (Atar, Aoujeft, Akjoujt; OIE), and RVF virus was eventually spread from southern to northern Mauritania by trade with infected animals that were brought there because of the unusually rich vegetation after the heavy rainfalls in the north. Furthermore, these heavy rainfalls caused an intensive increase in the mosquito population in that area facilitating the spread of RVFV (El Mamy et al., 2011). According to official numbers, there were 173 laboratory confirmed cases with 21 fatalities among camels and small ruminants (OIE). Although true numbers may have been much higher, 63 humans were clinically affected, 17 or 13 of these died depending on the source of information (El Mamy et al., 2011; Promedmail, 2010). In this study, we assayed serum samples collected in 2010 and 2011 from small ruminants, cattle and camels from various affected regions by molecular and serological tests to elucidate seroprevalence rates and transmission trails. Results highlight the importance of camelids as amplifying hosts during this epidemic. Materials and Methods Serum samples Between December 2010 and February 2011, serum samples were collected from sheep, goats, cattle and camels in RVF 32
affected regions of Mauritania. All samples were inactivated under BSL3 conditions with PBS–Tween (final concentration 0.5%) and by heating for 1 h at 56°C (Jansen Van Vuren and Paweska, 2010). RNA Isolation and quantitative real-time RT-PCR RNA was isolated from all serum samples using the QIAamp Viral RNA Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. All samples were tested with two different real-time RTPCR assays. Assay 1 targeted the L-segment at nucleotide (nt) position 2912–3001 (Bird et al., 2007). Assay 2 was a novel in-house real-time RT-PCR protocol, amplifying a fragment of about 160 nucleotides also on the L-segment (nucleotides 4447–4607 as calculated according to RVFV strain MP-12, GenBank accession number DQ375404.1). Primers and probes used for the amplification were f17: (5′-CCCAAGAGTCAACATCAGGGATGCA-3′) as forward primer, r151 (5′- TGGAGATGTTCGGAGGGTCTTCTG C-3′) as reverse primer and s95 (5′-FAM- AAGATRGTGA CCCTGGTTTGGGTTGCATCC-BHQ1-3′) as probe. Realtime RT-PCR was performed for both assays using the QuantiTectâ Probe RT-PCR Kit (Qiagen) as follows: 5 ll RNA, 20.0 pmol of each primer and 2.5 pmol of each probe were used for each reaction in a total volume of about 25 ll. Pure water instead of RNA was used as negative control in each run. Reaction conditions were 1 cycle at 50°C for 30 min (reverse transcription); 95°C for 15 min and 45 cycles at 95°C for 10 s, 55°C for 25 s and 72°C for 25 s. Quantification of viral load was carried out using an in-house synthetic RNA control (external calibrator) based on assay 1. For this purpose, the target region was inserted into vector pCR2.1 (MWG), which harbours a T7 promoter for in vitro transcription. The corresponding sequence was 5′-TAATGAAAATTCCTGAGACACATGGAC TCGCGCAGTGCCTGGATGACCCGGGCCAACCAATGT AGGGGCCTGTGTGGACTTGTGCAACATCAGATGATG CA AGGAAGT-3′. In contrast to the original sequence, a new specific target site was added, which encompassed 33 bp instead of 16 bp to enable binding of a second synthetic control probe. This new specific binding site was detected by the synthetic control probe (5′-CY5-CTC GCG CAG TGC CTG GAT GAC CCG GG-BHQ1-3′). Genetic analysis For genetic analysis of real-time RT-PCR positive samples, M-segment sequences were amplified using the Superscript III One-Step RT-PCR Kit (Qiagen) according to the manufacturer’s instructions. Primers were M-Sg forw (5′-ATG TATGTTTTATTAACAATT-3′), targeting nt position 21–41 and Gn-rev2 (5′-TTATGCTGATGCATATGAGA
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C-3′) targeting nt position 2073–2090 of the M-segment (according to RVFV strain MP-12; GenBank accession DQ380208.1). The following cycles were used as follows: one cycle at 55°C for 30 min and one cycle at 94°C for 2 min followed by 40 cycles at 94°C for 15 s, 60°C for 30 s and 68°C for 2.5 min followed by a final elongation step at 68°C for 10 min. Following electrophoretic separation, the purified genome fragments were sequenced. Sequence alignment and phylogenetic analysis were performed using Bioedit and MEGA 5 analysis software. RVFV competition ELISA All samples were tested with the ID Screen RVFV competition multispecies ELISA (ID Vet; Montpellier, France) according to the manufacturer’s instructions. Field samples blocking more than 60% of the RVFV detection antibodies compared with a negative control serum (i.e. OD field sample/OD negative control (S/N *100 in %) lower than 40) were qualified ‘positive’. Samples giving results between 40% and 50% were ‘inconclusive’, and results above 50% were considered ‘negative’. All sera were tested in duplicate. RVFV IgM capture ELISA Samples from small ruminants and cattle were also tested with the ID Screen Rift Valley Fever IgM capture ELISA (ID Vet; Montpellier, France) following the instruction manual. Samples producing OD field sample/OD positive control ratios (in%) above 50 were considered to be ‘positive’, those between 40 and 50 ‘inconclusive’ and samples with ratios lower than 40 ‘negative’. Indirect IgG DGn ELISA All samples from sheep and goats were tested in an inhouse indirect IgG ELISA based on bacterially expressed DGn protein. Production of recombinant antigen and the assay procedure was carried out as described previously (J€ackel et al., 2013). Maxisorb immunoplates (Nunc, Roskilde, Denmark) were coated with recombinant RVFV DGn protein, diluted in 0.05 M carbonate–bicarbonate buffer (pH 9.6) to a concentration of 2 lg/ml (100 ll diluted antigen per well). After incubating the plates overnight at 4°C, they were washed three times with 300 ll washing buffer (phosphate-buffered saline (PBS) pH 7.2; 0.1% Tween 20), followed by a blocking step with 200 ll/ well 10% skim milk powder (DIFCOTM) diluted in PBS for 1 h at 37°C in a moist chamber. Polyclonal hyperimmune rabbit serum as positive control was diluted 1 : 20 000 in PBS containing 2% skim milk powder (dilution buffer), and the negative control (negative goat serum) and Mau-
Rift Valley Fever Outbreak in Mauritania
ritanian serum samples were diluted 1 : 25 in dilution buffer. The positive control was tested in quadruplicate whereas the negative control, conjugate control (2% skim milk) and each sample were tested in duplicate (100 ll per well). Plates were again incubated at 37°C for 1 h in a moist chamber, followed by a further washing step as described above. Horseradish peroxidase-conjugated Protein G (Calbiochem) was diluted 1 : 5000 in dilution buffer, and 100 ll were then added to each well. After another 1 h incubation step, plates were washed again and 100 ll substrate (2,2′-azino di-ethylbenzothiazoline sulphonic acid (ABTS); Roche, Mannheim, Germany) was added to each well. Reactions were run for 30 min at room temperature in the dark, stopped by addition of 1% sodium dodecyl sulphate and the optical density (OD value) at 405 nm was determined. Results were expressed as percentages of the positive control serum (PP value): (mean OD of duplicate test serum/median OD of quadruplicate positive control) 9 100. All samples with PP values lower than the cut-off were considered to be ‘negative’, sera with higher PP values ‘positive’. Indirect immunofluorescence assay All sera were screened for anti-RVFV IgG antibodies by indirect immunofluorescence assay (IIFA) using a modified commercial RVFV IIFA slide test kit (Euroimmun, L€ ubeck, Germany). Slides containing a mixture of infected and non-infected Vero E6 cells on one field (positive field) and non-infected Vero E6 cells on a negative control field were used. Sheep, goat and cattle sera were diluted 1 : 100 with sampling buffer, and 25 ll of the diluted samples were applied to a biochip and incubated for 30 min at room temperature. After a first washing step for 10 min with phosphate-buffered saline (PBS), pH 7.2; 0.2% Tween 20, 25 ll of donkey anti-sheep IgG Cy3 (Indocarbocyanin)-labelled antibodies (Dianova, Hamburg, Germany) for small ruminants and goat anti-bovine IgG Cy3-labelled antibodies (Dianova) diluted 1 : 200 in PBS containing 0.2% Tween were applied. For all small ruminant samples, anti-sheep secondary antibody was used. After a second incubation step at room temperature in the dark for 30 min, the slides were washed again for 10 min, dried and covered with sampling buffer and cover plates. Slides were then examined on a fluorescence microscope (Nikon, Tokyo, Japan). A modification was necessary for testing camel sera as no fluorescence-labelled secondary antibody to camelid IgG was available. Therefore, a polyclonal anti-camel antiserum produced in rabbits (Bethyl laboratories, Montgomery, TX, USA) was used instead (dilution 1 : 100 in sampling buffer) followed by a Cy3 conjugated goat anti-rabbit antibody (Dianova) diluted 1 : 800 in PBS with 0.2% Tween.
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different real-time RT-PCR assays. Specific RVFV RNA could be detected in both assays in four camels (Fig. 2a). All real-time RT-PCR positive samples were collected in the central Mauritanian region of Adrar in the department of Aoujeft. Ct-values ranged from 24.9 to 35.71 (Fig. 2a). The viral load was highest in sample # 8 with 143 270 copies per reaction and Ct-values of 24.9 (assay 1) and 26.22 (assay 2), followed by sample # 9 with more than 7000 copies per reaction and Ct-values of 29.36 (assay 1) and 29.95
Results Quantitative real-time RT-PCR and genetic analysis From December 2010 until February 2011, 163 samples from small ruminants (sheep and goats), cattle and camels were collected in nine different Mauritanian regions throughout the country (Figs 1a and b). The collection included serum samples from 93 small ruminants, 62 camels and eight bovines. All samples were tested with two
(b)
(a)
region
number of sera
small ruminants
camel
cattle
Inchiri/Akjoujt
40
40
-
-
Adrar/Aoujeft
38
28
10
-
Nouadibou
39
-
39
-
Brakna
3
-
3
-
Nouakchott
8
7
-
1
Assaba/Kankossa
1
-
-
1
Gorgol/Mbout
12
2
10
-
Trarza/Ouad Naga
8
2
-
6
Erch
14
14
-
-
Total
163
93
62
8
Fig. 1. (a) Outbreak map of Rift zalley fever virus in Mauritania in 2010. Three regions in northern parts of the country were affected by the virus. (b) Sample origin and numbers. In total, 163 samples were collected from small ruminants (n = 93), camelids (n = 62) and cattle (n = 8) from nine different regions of Mauritania.
(a)
sample
species
region
assay 1
cop/rx
assay 2
sample 8
camel
sample 9
camel
Adrar/Aoujeft
24,9
143270
26,22
Adrar/Aoujeft
29,36
7181
29,95
p 124 sample
camel
Adrar/Aoujeft
34,13
103
35,71
sample 125
camel
Adrar/Aoujeft
34,74
67,9
33,78
(b) 96 DQ380186 OS-1 Mauritania 1987 human 63 54
DQ380184 OS-3 Mauritania 1987 human JQ974833 Mauritania camel NC_014396 Z H548 Egypt 1977 DQ380197 Saudi Arabia 2000 human NC_005220 S23 Uukuniemi
0.1
Bootstrap consensus tree (1000 replicates) Fig. 2. (a) Real-time RT-PCR results from four positive camel samples. Specific viral RNA from Rift Valley fever virus (RVFV) could be detected with two different real-time RT-PCR assays. (b) Phylogenetic analysis of an 859 nucleotide sequence from the M-segment detected in a Mauritanian camel (sample number 8). Sequences were analysed with 1000 replicate bootstrap values using Kimura 2 parameter model. Numbers indicate the percentage of 1000 bootstrap replicates that support each labelled node. For each sequence, the GenBank accession number, strain designation and strain origin are provided. Analysis was supported by the use of Uukuniemi strain S23 as out-group.
34
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(assay 2). The viral load of samples # 124 and # 125 with Ct-values above 33 in both assays was lower with 103 and 68 copies per reaction, respectively. From sample # 8, a partial M-segment sequence of about 859 nucleotides was generated and sequenced [GenBank accession JQ974833; nt position 66–924, according to RVFV strain ZH-548 (GenBank accession NC 014396.1)]. A phylogenetic analysis was performed using sequences from Mauritania 1987, Egypt 1977 and Saudi Arabia 2000. The newly generated camel sequence was closely associated with viruses from the Mauritanian outbreak in 1987 (Fig. 2b). Only three amino acid differences were found between the isolates from 1987 to 2010. However, there were only four different amino acids compared with RVFV strain ZH-548 and seven different amino acids compared with the Saudi Arabian isolate. As all sequences shared a nucleotide homology between 94% and 95%, a more detailed genetic analysis cannot be carried out. Partial sequences derived from the Mauritanian outbreak in 2003 could not be included in the phylogenetic analysis as they overlapped just in a short section (nt 823–924) with the newly generated camel sequence. RVFV competition ELISA and IgM capture ELISA A total of 93 samples (small ruminants n = 64; cattle n = 1; camel n = 28) were positive in the ID Screen RVFV competition multispecies ELISA, which detects IgG as well as IgM antibodies. Four samples gave inconclusive results, and 66 samples were negative (Table 1A; columns 2–4). Overall seroprevalence rates were found in Adrar/Aoujeft (87%), Inchiri/Akjoujt (78%), Nouadibou (41%) and Erch (57%; Table 1B, column 3). The seroprevalence rate among small ruminants was 69% compared with 45% in camels (Table 1A, column 5). Just one bovine sample from eight tested sera was positive in the competition ELISA (Table 1A, column 2). The highest RVF antibody seroprevalence in small ruminants was found in Adrar/Aoujeft (86%) and Inchiri/Akjoujt (78%), whereas affected camels originated mainly from Adrar/Aoujeft and Nouadibou regions (Table 2). A total of 49 samples (all small ruminants) were positive in the ID Screen Rift Valley Fever IgM capture ELISA giving evidence for recent infections by the virus, four samples had a doubtful result and in 48 samples (small ruminants n = 40; cattle n = 8), and no RVFV-specific IgM antibodies were detected (Table 1A, columns 6–8). The highest number of positive small ruminants was found in Inchiri/ Akjoujt with 63%, followed by Adrar/Aoujeft and Erch, both with 57% IgM-positive animals (Table 1B, column 5). In other regions, no further IgM-positive animals were found. All bovine samples were IgM negative (Table 1A, column 9). Camels could not be tested for the presence of
Rift Valley Fever Outbreak in Mauritania
IgM antibodies as the commercial IgM capture ELISA is not functioning in camelids. In-house indirect IgG DGn ELISA All small ruminant samples were also analysed with an in-house indirect IgG DGn-based ELISA. The results are displayed in Table 1A (column 10–12) and Table 1B (columns 6–7). In total, 51 samples were positive with a PP value above 20 752. No RVFV-specific IgG antibodies were detected in the remaining 42 samples. The seroprevalence was 55% with the highest prevalence of IgG antibodies in Adrar/Aoujeft (79%) followed by Inchiri/Akjoujt (60%), Erch (29%) and Nouakchott (14%). There were no IgGpositive animals in Trarza and Gorgol. Comparison of all three ELISA assays revealed additional results for the infection status of small ruminants (Table 3): 36 of the 51 DGn ELISA-positive samples were also positive with both commercial ELISAs. Eleven samples showed evidence of a past infection as they were just positive with the IgG DGn-based ELISA and with the ID Screen competition ELISA. In 13 animals, a recent infection could be shown as they were just positive with the two commercial ELISAs, both detecting IgM antibodies. Indirect immunofluorescence assay All samples were eventually tested in an IIFA for the presence of RVFV-specific IgG antibodies (using speciesspecific secondary antibodies). As shown in Fig. 3, in sera of all four species, RVF virus antibodies were detected [see Figs 3a (sheep), b (goat), c (camel) and d (cattle)]. With regard to small ruminants, 57 sera showed positive staining; eight had a doubtful result and no RVFV-specific IgG antibodies were detected in 28 samples (Table 1A; columns 13–16). Just two of eight cattle samples were positive in indirect immunofluorescence; in the remaining six samples, no IgG antibodies could be detected. 27 camels were positive, 11 sera had a doubtful result, whereas in 24 samples no antibodies were found (Table 1A; columns 13–16). Prevalence was highest in Adrar/Aoujeft with 82% (Table 1B, columns 8–9). In Inchiri/Akjoujt, the seroprevalence was 75%, for Erch it was lower (43%). In Gorgol/Mbout and Brakna, the seroprevalence in the immunofluorescence assay was 33%, followed by Nouadibou with 31% and Trarza/Ouad Naga where two of eight tested animals were positive (25%). In Nouakchott and Assaba/Kankossa, no IIFApositive animals were found (Table 1B, columns 8–9). Discussion In October/November 2010, a massive outbreak of Rift Valley fever occurred in northern Mauritania after unusually
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36 32 7 66
27
Negative
45 13 57
69
Prevalence (%)
31/40 33/38 16/39 1/3 1/8 0/1 2/12 1/8 8/14 93/163
Inchiri/Akjoujt Adrar/Aoujeft Nouadibou Brakna Nouakchott Assaba/Kankossa Gorgol/Mbout Trarza/Ouad Naga Erch Total
78 87 41 33 13 0 17 13 57 57
Prevalence (%)
n.t. 0 49
49 n.t. 0 4
4
Doubtful
n.t. 8 48
40
Negative
n.t. n.t. 51
– 0 59
25/40 16/28 n.t. n.t. 0/8 0/1 0/2 0/8 8/14 49/101
Positive/total 63 57 n.t. n.t. 0 0 0 0 57 49
Prevalence (%)
n.t. n.t. 42
42
Negative
– – 55
55
Prevalence (%)
24/40 22/28 n.t. n.t. 1/7 n.t. 0/2 0/2 4/14 51/93
Positive/total
27 2 86
57
Positive
60 79 n.t. n.t. 14 n.t. 0 0 29 55
24 6 58
28
Negative
a
44 25 53
61
Prevalence (%)
30/40 31/38 12/39 1/3 0/8 0/1 4/12 2/8 6/14 86/163
Positive/total
75 82 31 33 0 0 33 25 43 53
Prevalence (%)
Indirect immunofluorescence assay
11 0 19
8
Doubtful
Indirect immunofluorescence assay
Prevalence (%)
Indirect IgG Gn ELISAb
51
Positive
Indirect IgG Gn ELISAb
53
Prevalence (%)
ID screen IgM capture ELISAa
Positive
ID screen IgM capture ELISAa
The number of positive, negative and doubtful samples for each assay and species as well as the resulting seroprevalence is shown. The percentages were rounded off to whole numbers. Camels not tested. b Camels and cattle not tested.
Positive/total
Region
ID screen competition ELISA
2 0 4
28 1 93
(B)
2
64
Small ruminants Camel Cattle Total
Doubtful
Positive
ID screen competition ELISA
Species
(A)
Table 1. Serological analysis of Mauritanian serum samples from small ruminants, cattle and camels with three different ELISAs (ID Vet ID Screen Rift Valley Fever competition ELISA (multispecies); ID Vet ID Screen Rift Valley Fever IgM capture ELISA and in-house indirect IgG DGn ELISA) and indirect immunofluorescence assay (IIFA). Camels were only tested with the ID VET competition ELISA and IIFA, cattle samples were not tested with the in-house indirect IgG ELISA. (A) Results of the four assays grouped into species. (B) Results of the four assays with regard to geographic distribution
Rift Valley Fever Outbreak in Mauritania S. J€ackel et al.
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Table 2. Seroprevalence of IgG/IgM antibodies in different Mauritanian regions determined with the ID VET competition ELISA subdivided into species. Small ruminants
Cattle
Region
Positive/ total
%
Inchiri/Akjoujt Adrar/Aoujeft Nouadibou Brakna Nouakchott Assaba/Kankossa Gorgol/Mbout Trarza/Ouad Naga Erch Total
31/40 24/28 – – 1/7 – 0/2 0/2 8/14 64/93
78 86 – – 14 – 0 0 57 69
Camels
Positive/ total
%
– – – – 0/1 0/1 – 1/6
– – – – 0 0 – 17
1/8
13
Positive/ total
%
– 9/10 16/39 1/3 – – 2/10 – – 28/62
– 90 41 33 – – 20 – – 45
long and heavy rainfalls. Humans, livestock and camels were affected (El Mamy et al., 2011). In the here presented follow-up study, we assayed field samples collected in the late time period of this epidemic (December 2010 to February 2011) to monitor whether RVFV still circulated in animals at this time. As all samples in this as well as in an earlier study (El Mamy et al., 2011) were regular submissions to the diagnostic laboratory, they do not represent a random selection, that is, results need to be interpreted carefully. In our study, sera from livestock were tested for viremia by real-time qRT-PCR and run in two commercially available ELISAs. Moreover, small ruminant sera were also tested in a newly developed indirect IgG ELISA (J€ackel et al., 2013). Our results show that the overall seroprevalence rate for IgG/IgM antibodies (as measured by ID VET competition ELISA) in livestock animals and camels increased from 37% to 57% when compared with recently published
Table 3. Identification of the serological status of small ruminants by comparing three different ELISAs. Samples with a doubtful result in competition ELISA and/or IgM capture ELISA are not shown
ELISA result
Competition ELISA positive IgM capture ELISA postive Indirect IgG Gn ELISA positive
Competition ELISA positive IgM capture ELISA postive Indirect IgG Gn ELISA negative
Competition ELISA positive IgM capture ELISA negative Indirect IgG Gn ELISA positive
Competition ELISA positive IgM capture ELISA negative Indirect IgG Gn ELISA negative
Sample number
36 (IgM and IgG)
13 (IgM only)
11 (IgG only)
27 (negative)
(a)
(b)
Mau123 (c)
Mau10 (d)
Mau7
Mau146
€beck, Germany) Fig. 3. Indirect immunofluorescence assay (IIFA) using a commercial Rift Valley fever virus (RVFV) IIFA slide test kit (Euroimmun, Lu with slight modifications for testing ruminant and camel serum samples. (a) sheep, (b) goat, (c) camel, (d) cattle.
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results generated from sera sampled in September and October 2010 (El Mamy et al., 2011). In particular, the seroprevalence in small ruminants rose from 43% to 69% over this time. Furthermore, there was an increase in the number of IgM-positive animals (53% compared with 42%). Used in combination, the three assays demonstrated that only 11 of 93 small ruminants showed exclusively IgG antibodies, which points to a recent virus exposure of the majority of sampled animals. The IIFA, which was originally designed for testing human sera, was adapted to animal sera and yielded results, which clearly correlated with the ELISA results. Camels played an important role in RVFV epidemics in Kenya (Scott et al., 1963; Davies et al., 1985), Nigeria (Davies et al., 1985), Sudan (Eisa, 1984), Egypt (Davies et al., 1985; Ahmed Kamal, 2011) and Morocco (El-Harrak et al., 2011). During the first RVF outbreak in Egypt, camels which were moved from northern Sudan to the southern parts of Egypt may have spread the virus (Ahmed Kamal, 2011). In Morocco, El-Harrak et al. found a seroprevalence of 15% among camels in 2009 and suspected animal movements from south-eastern parts of the Sahara desert to the north-west as reasons. After an earlier outbreak in Mauritania in 1987, camels carried RVF antibodies at a rate of about 33% (Saluzzo et al., 1987). In 1998, Nabeth et al. found one IgM-positive sample of 39 tested camels. Camels also played a very important role in the RVF outbreak in Mauritania in 2010, and numerous animals showed clinical signs. Rift Valley fever virus RNA was detected in seven animals by real-time RT-PCR, and virus was isolated from four of these samples (El Mamy et al., 2011). The overall seroprevalence rate for IgG/IgM antibodies in camels was again about 33%. In the here described study on late epidemic sera, a seroprevalence rate of about 45% was found in camels, which underlines their central role during the latest Mauritanian outbreak. Moreover, using two different quantitative real-time RT-PCR assays, we detected RVFV RNA in four camelid samples and a partial sequence of the M-segment was obtained from one animal (GenBank accession number: JQ974833). To our knowledge, this is the first published camel-derived RVFV sequence apart from sequences originating from ticks feeding on a camel in 1979 (accession numbers: HQ403564.1; HQ403562.1; HQ403563.1). The phylogenetic analysis showed that this sequence is closely associated with those of viruses from the Mauritanian outbreak in 1987. However, due to the low variability and a high nucleotide homology of >95% to other virus sequences, further genetic analysis of this fragment was not possible. In conclusion, we could show that the RVF virus was circulating in Mauritania after the initial outbreak in October 2010 at least until January 2011, when the epidemic was officially declared to be over, since two camels were still viremic 38
then. Furthermore, our results highlight the important role of small ruminants during the infection cycle of RVFV. Acknowledgements We would like to acknowledge Dr. Modou Moustapha L^ o, L’Institut senegalais de recherches agricoles (ISRA) / Laboratoire National de l’Elevage et de Recherches Veterinaires (LNERV), Dakar, Senegal, Dr. Anne Balkema-Buschmann, Friedrich-Loeffler-Institut and Dr. Hermann Unger, Joint FAO/IAEA Division, Vienna, Austria, for their scientific and technical support of this study. This study was partially funded by EU grant FP7-261504 EDENext and is catalogued by the EDENext Steering Committee as EDENext159 (\http://www.edenext.eu) as well as by the FP7 NADIR project. The contents of this publication are the sole responsibility of the authors and do not necessarily reflect the views of the European Commission. Conflicts of interest The authors have no conflicts of interest to declare. References Ahmed Kamal, S., 2011: Observations on rift valley fever virus and vaccines in Egypt. Virol. J. 8, 532. Bird, B. H., D. A. Bawiec, T. G. Ksiazek, T. R. Shoemaker, and S. T. Nichol, 2007: Highly sensitive and broadly reactive quantitative reverse transcription-PCR assay for high-throughput detection of Rift Valley fever virus. J. Clin. Microbiol. 45, 3506–3513. Bird, B. H., T. G. Ksiazek, S. T. Nichol, and N. J. MacLachlan, 2009: Rift Valley fever virus. J. Am. Vet. Med. Assoc. 234, 883–893. Bouloy, M., and F. Weber, 2010: Molecular biology of rift valley fever virus. Open Virol. J. 4, 8–14. Chevalier, V., T. Rakotondrafara, M. Jourdan, J. M. Heraud, H. R. Andriamanivo, B. Durand, J. Ravaomanana, P. E. Rollin, and R. Rakotondravao, 2011: An unexpected recurrent transmission of Rift Valley fever virus in cattle in a temperate and mountainous area of Madagascar. PLoS Negl. Trop. Dis. 5, e1423. Daubney, R., J. R. Hudson, and P. C. Garnham, 1931: Enzootic hepatitis or rift valley fever. An undescribed virus disease of sheep, cattle and man from East Africa. J. Pathol. Bacteriol. 34, 545–579. Davies, F. G., J. Koros, and H. Mbugua, 1985: Rift Valley fever in Kenya: the presence of antibody to the virus in camels (Camelus dromedarius). J. Hyg. (Lond) 94, 241–244. Digoutte, J. P., and C. J. Peters, 1989: General aspects of the 1987 Rift Valley fever epidemic in Mauritania. Res. Virol. 140, 27–30. Eisa, M., 1984: Preliminary survey of domestic animals of the Sudan for precipitating antibodies to Rift Valley fever virus. J. Hyg. (Lond) 93, 629–637.
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El Mamy, A. B., M. O. Baba, Y. Barry, K. Isselmou, M. L. Dia, M. O. El Kory, M. Diop, M. M. Lo, Y. Thiongane, M. Bengoumi, L. Puech, L. Plee, F. Claes, S. de la Roque, and B. Doumbia, 2011: Unexpected Rift Valley fever outbreak, northern Mauritania. Emerg. Infect. Dis. 17, 1894–1896. El-Harrak, M., R. Martın-Folgar, F. Llorente, P. FernandezPacheco, A. Brun, J. Figuerola, and M. A. Jimenez-Clavero, 2011: Rift Valley and West Nile virus antibodies in camels, North Africa. Emerg. Infect. Dis. 17, 2372–2374. Faye, O., M. Diallo, D. Diop, O. E. Bezeid, H. B^a, M. Niang, I. Dia, S. A. Mohamed, K. Ndiaye, D. Diallo, P. O. Ly, B. Diallo, P. Nabeth, F. Simon, B. L^ o, and O. M. Diop, 2007: Rift Valley fever outbreak with East-Central African virus lineage in Mauritania, 2003. Emerg. Infect. Dis. 13, 1016–1023. Gad, A. M., F. M. Feinsod, I. H. Allam, M. Eisa, A. N. Hassan, B. A. Soliman, S. el Said, and A. J. Saah, 1986: A possible route for the introduction of Rift Valley fever virus into Egypt during 1977. J. Trop. Med. Hyg. 89, 233–236. Hassan, O. A., C. Ahlm, R. Sang, and M. Evander, 2011: The 2007 Rift Valley fever outbreak in Sudan. PLoS Negl. Trop. Dis. 5, e1229. Hightower, A., C. Kinkade, P. M. Nguku, A. Anyangu, D. Mutonga, J. Omolo, M. K. Njenga, D. R. Feikin, D. Schnabel, M. Ombok, and R. F. Breiman, 2012: Relationship of climate, geography, and geology to the incidence of Rift Valley fever in Kenya during the 2006–2007 outbreak. Am. J. Trop. Med. Hyg. 86, 373–380. J€ackel, S., M. Eiden, A. Balkema-Buschmann, M. Ziller, P. Jansen van Vuren, J. T. Paweska, and M. H. Groschup, 2013: A novel indirect ELISA based on glycoprotein Gn for the detection of IgG antibodies against Rift Valley fever virus in small ruminants. Res. Vet. Sci. in press, doi: 0.1016/j.rvsc.2013.04.015. [Epub ahead of print]. Jansen Van Vuren, P., and J. T. Paweska, 2010: Comparison of enzyme-linked immunosorbent assay-based techniques for the detection of antibody to Rift Valley fever virus in thermochemically inactivated sheep sera. Vector Borne Zoonotic Dis. 10, 697–699. Jeanmaire, E. M., R. Rabenarivahiny, M. Biarmann, L. Rabibisoa, L. F. Ravaomanana, T. Randriamparany, S. F. Andriamandimby, C. S. Diaw, P. Fenozara, S. de la Roque, and J. M. Reynes, 2011: Prevalence of Rift Valley fever infection in ruminants in Madagascar after the 2008 outbreak. Vector Borne Zoonotic Dis. 11, 395–402. Jost, C. C., S. Nzietchueng, S. Kihu, B. Bett, G. Njogu, E. S. Swai, and J. C. Mariner, 2010: Epidemiological assessment of the Rift Valley fever outbreak in Kenya and Tanzania in 2006 and 2007. Am. J. Trop. Med. Hyg. 83(2 Suppl.), 65–72. Jouan, A., B. Le Guenno, J. P. Digoutte, B. Philippe, O. Riou, and F. Adam, 1988: An RVF epidemic in southern Mauritania. Ann. Inst. Pasteur Virol. 139, 307–308. Ksiazek, T. G., A. Jouan, J. M. Meegan, B. Le Guenno, M. L. Wilson, C. J. Peters, J. P. Digouette, M. Guillaud, N. O. Merzoug, and E. M. Touray, 1989: Rift Valley fever among domestic animals in the recent West African outbreak. Res. Virol. 140, 67–77.
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Moutailler, S., G. Krida, F. Schaffner, M. Vazeille, and A. B. Failloux, 2008: Potential vectors of Rift Valley fever virus in the Mediterranean region. Vector Borne Zoonotic Dis. 8, 749– 753. Munyua, P., R. M. Murithi, S. Wainwright, J. Githinji, A. Hightower, D. Mutonga, J. Macharia, P. M. Ithondeka, J. Musaa, R. F. Breiman, P. Bloland, and M. K. Njenga, 2010: Rift Valley fever outbreak in livestock in Kenya, 2006–2007. Am. J. Trop. Med. Hyg. 83(2 Suppl.), 58–64. Nabeth, P., Y. Kane, M. O. Abdalahi, M. Diallo, K. Ndiaye, K. Ba, F. Schneegans, A. A. Sall, and C. Mathiot, 2001: Rift Valley fever outbreak, Mauritania, 1998: seroepidemiologic, virologic, entomologic, and zoologic investigations. Emerg. Infect. Dis. 7, 1052–1054. Nguku, P. M., S. K. Sharif, D. Mutonga, S. Amwayi, J. Omolo, O. Mohammed, E. C. Farnon, L. H. Gould, E. Lederman, C. Rao, R. Sang, D. Schnabel, D. R. Feikin, A. Hightower, M. K. Njenga, and R. F. Breiman, 2010: An investigation of a major outbreak of Rift Valley fever in Kenya: 2006–2007. Am. J. Trop. Med. Hyg. 83(2 Suppl.), 5–13. OIE 2011: World Animal Health Information Database (WAHID). Available at \http://web.oie.int/wahis/public.php? WAHIDPHPSESSID=954bacd41c95f0ac7b7cdf6965375e39& page=disease_immediate_summary&disease_type=Terrestrial&disease_id=8 (accessed March 8, 2011). Pepin, M., M. Bouloy, B. H. Bird, A. Kemp, and J. T. Paweska, 2010: Rift Valley fever virus (Bunyaviridae: Phlebovirus): an update on pathogenesis, molecular epidemiology, vectors, diagnostics and prevention. Vet. Res. 41, 1–40. Promedmail 2010: Published Date: 2010-12-02 19:00:05. Subject: PRO/AH/EDR> Rift Valley fever, livestock, human – Mauritania. Archive Number: 20101202.4332 Promedmail 2012: Published Date: 2012-10-05 19:34:27. Subject: PRO/AH/EDR> Rift Valley fever – Mauritania (02): fatalities. Archive Number: 20121005.1326899 Saluzzo, J. F., J. P. Digoutte, C. Chartier, D. Martinez, and R. Bada, 1987: Focus of Rift Valley fever virus transmission in southern Mauritania. Lancet 1, 504. Saluzzo, J. F., G. W. Jr Anderson, L. A. Hodgson, and J. P. Digoutte, 1989: Antigenic and biological properties of Rift Valley fever virus isolated during the 1987 Mauritanian epidemic. Res. Virol. 140, 155–164. Scott, G. R., W. Coackley, R. W. Roach, and N. R. Cowdy, 1963: Rift Valley fever in camels. J. Pathol. Bacteriol. 86, 229–231. Seufi, A. M., and F. H. Galal, 2007: Role of Culex and Anopheles mosquito species as potential vectors of rift valley fever virus in Sudan outbreak. BMC Infect. Dis. 10, 65. Swai, E. S., and L. Schoonman, 2009: Prevalence of Rift Valley fever immunoglobulin G antibody in various occupational groups before the 2007 outbreak in Tanzania. Vector Borne Zoonotic Dis. 9, 579–582. Zeller, H. G., A. J. Akakpo, and M. M. Ba, 1995: Rift Valley fever epizootic in small ruminants in southern Mauritania (October 1993): risk of extensive outbreaks. Ann. Soc. Belg. Med. Trop. 75, 135–140.
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ORIGINAL ARTICLE
Molecular and Serological Studies on the Rift Valley Fever Outbreak in Mauritania in 2010 €ckel1, M. Eiden1, B. O. EL Mamy2, K. Isselmou2, A. Vina-Rodriguez1, B. Doumbia3 and S. Ja M. H. Groschup1 1 2 3
Institute for Novel and Emerging Infectious Diseases, Friedrich-Loeffler-Institut, Greifswald - Insel Riems, Germany Service de Pathologie Infectieuses, Centre National de l’Elevage et de Recherches Veterinaires (CNERV), Nouakchott, Mauritanie Minist ere du d eveloppement rural, Nouakchott, Mauritania
Keywords: Rift Valley fever virus; small ruminants; camels; Mauritania Correspondence: M. H. Groschup. Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of Novel and Emerging Infectious €dufer 10, 17493 Greifswald - Isle Diseases, Su of Riems, Germany. Tel.: +49 38351 71163; Fax: +49 38351 71191; E-mail: martin. [email protected] Received for publication November 15, 2012 doi:10.1111/tbed.12142
Summary Rift Valley fever virus (RVFV) is a vector-borne RNA virus affecting humans, livestock and wildlife. In October/November 2010, after a period of unusually heavy rainfall, a Rift Valley fever outbreak occurred in northern Mauritania causing clinical cases in cattle, sheep, goats and camels, 21 of which were of lethal outcome. The aim of this study was to obtain further information on the continuation of RVF virus activity and spread in animal species in Mauritania after this outbreak. We therefore tested sera from small ruminants, cattle and camels for the presence of viral RNA and antibodies against RVFV. These sera were collected in different parts of the country from December 2010 to February 2011 and tested with three different ELISAs and an indirect immunofluorescence assay. The results show a high seroprevalence of RVFV IgM and IgG antibodies of about 57% in all animals investigated. Moreover, in four camel sera, viral RNA was detected emphasizing the important role camels played during the latest RVF outbreak in Mauritania. The study demonstrates the continuous spread of RVFV in Mauritania after initial emergence and highlights the potential role of small ruminants and camels in virus dissemination.
Introduction Rift Valley fever virus (RVFV) is a zoonotic arbovirus affecting humans and a wide range of vertebrate hosts such as small ruminants, cattle, camels and wild ruminants. Rift Valley fever virus was first discovered in 1931 in Kenya (Great Rift Valley) in a huge outbreak, which claimed the lives of more than 4500 lambs and ewes (Daubney et al., 1931; referenced in Bird et al., 2009). Rift Valley fever virus is a single-stranded RNA virus belonging to the family Bunyaviridae; genus Phlebovirus. Its genome consists of three segments, the M (medium)- and L (large)-segment are of negative orientation whereas the S (small)-segment has ambisense polarity. The S-segment encodes the nucleocapsid protein (N-protein) and a non-structural protein Nss. The M-segment encodes the two glycoproteins Gn and Gc and also includes the genetic information for a nonstructural protein Nsm, whereas the L-segment codes for the RNA-dependent RNA-polymerase (Pepin et al., 2010).
Numerous mosquito species are considered to be competent RVFV transmission vectors, several of them can also be found in Europe (Moutailler et al., 2008; Pepin et al., 2010). Rift Valley fever virus infections in humans usually lead to a self-limiting febrile illness. However, in 1–2% of these infections, humans develop retinitis, encephalitis, haemorrhagic fever and acute hepatitis, sometimes with lethal outcome. Morbidity and mortality in livestock are much higher, as infections are characterized by high mortality rates of up to 100% among newborn lambs and by high abortion rates, especially among pregnant sheep and goats, the so-called ‘abortion storms’. Rift Valley fever virus transmission to humans can occur by direct contact with infected body fluids, tissues and/or excretions or via mosquito bites (Bird et al., 2009; Bouloy and Weber, 2010). The infection risk is especially high after periods of long and heavy rainfalls, as mosquitoes in suddenly flooded puddles and river beds hatch from infected eggs in which
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the virus can survive over many years. Blood-feeding female mosquitoes transmit RVFV to animals, which act as first amplification host and eventually infect humans (Bird et al., 2009). Until 1977, RVF outbreaks were restricted to sub-Saharan Africa, but more recently cases have also been found in Egypt (Ahmed Kamal, 2011) and Sudan (Gad et al., 1986). The first outbreak outside of the African continent was in 2000/2001 in Saudi Arabia and Yemen. Several outbreaks have been reported in different African countries since 2006: Sudan in 2007 (Seufi and Galal, 2007;; Hassan et al., 2011), Kenya in 2006/2007 (Munyua et al., 2010; Nguku et al., 2010; Hightower et al., 2012), Tanzania in 2007 (Swai and Schoonman, 2009; Jost et al., 2010) and Madagascar in 2008/9 (Chevalier et al., 2011; Jeanmaire et al., 2011). In 2010, more outbreaks were found in South Africa, Namibia, Saudi Arabia, Botswana and Mauritania (OIE; WAHID interface) with a total of more than 14 500 clinical cases and over 9000 dead livestock and wild animals. In Mauritania and Namibia, outbreaks resumed in early 2011, whereas cases in South Africa continued throughout the year. The first RVF cases were seen in Mauritania in 1987 when 220 humans died from the disease (Saluzzo et al., 1987, 1989; Jouan et al., 1988; Digoutte and Peters, 1989; Ksiazek et al., 1989). More RVF epidemics occurred subsequently in 1993 (Zeller et al., 1995), 1998 (Nabeth et al., 2001), 2003 (Faye et al., 2007), 2010 (El Mamy et al., 2011) and 2012 (Promedmail, 2012). Outbreaks in 2010 were concentrated on three regions (Atar, Aoujeft, Akjoujt; OIE), and RVF virus was eventually spread from southern to northern Mauritania by trade with infected animals that were brought there because of the unusually rich vegetation after the heavy rainfalls in the north. Furthermore, these heavy rainfalls caused an intensive increase in the mosquito population in that area facilitating the spread of RVFV (El Mamy et al., 2011). According to official numbers, there were 173 laboratory confirmed cases with 21 fatalities among camels and small ruminants (OIE). Although true numbers may have been much higher, 63 humans were clinically affected, 17 or 13 of these died depending on the source of information (El Mamy et al., 2011; Promedmail, 2010). In this study, we assayed serum samples collected in 2010 and 2011 from small ruminants, cattle and camels from various affected regions by molecular and serological tests to elucidate seroprevalence rates and transmission trails. Results highlight the importance of camelids as amplifying hosts during this epidemic. Materials and Methods Serum samples Between December 2010 and February 2011, serum samples were collected from sheep, goats, cattle and camels in RVF 32
affected regions of Mauritania. All samples were inactivated under BSL3 conditions with PBS–Tween (final concentration 0.5%) and by heating for 1 h at 56°C (Jansen Van Vuren and Paweska, 2010). RNA Isolation and quantitative real-time RT-PCR RNA was isolated from all serum samples using the QIAamp Viral RNA Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. All samples were tested with two different real-time RTPCR assays. Assay 1 targeted the L-segment at nucleotide (nt) position 2912–3001 (Bird et al., 2007). Assay 2 was a novel in-house real-time RT-PCR protocol, amplifying a fragment of about 160 nucleotides also on the L-segment (nucleotides 4447–4607 as calculated according to RVFV strain MP-12, GenBank accession number DQ375404.1). Primers and probes used for the amplification were f17: (5′-CCCAAGAGTCAACATCAGGGATGCA-3′) as forward primer, r151 (5′- TGGAGATGTTCGGAGGGTCTTCTG C-3′) as reverse primer and s95 (5′-FAM- AAGATRGTGA CCCTGGTTTGGGTTGCATCC-BHQ1-3′) as probe. Realtime RT-PCR was performed for both assays using the QuantiTectâ Probe RT-PCR Kit (Qiagen) as follows: 5 ll RNA, 20.0 pmol of each primer and 2.5 pmol of each probe were used for each reaction in a total volume of about 25 ll. Pure water instead of RNA was used as negative control in each run. Reaction conditions were 1 cycle at 50°C for 30 min (reverse transcription); 95°C for 15 min and 45 cycles at 95°C for 10 s, 55°C for 25 s and 72°C for 25 s. Quantification of viral load was carried out using an in-house synthetic RNA control (external calibrator) based on assay 1. For this purpose, the target region was inserted into vector pCR2.1 (MWG), which harbours a T7 promoter for in vitro transcription. The corresponding sequence was 5′-TAATGAAAATTCCTGAGACACATGGAC TCGCGCAGTGCCTGGATGACCCGGGCCAACCAATGT AGGGGCCTGTGTGGACTTGTGCAACATCAGATGATG CA AGGAAGT-3′. In contrast to the original sequence, a new specific target site was added, which encompassed 33 bp instead of 16 bp to enable binding of a second synthetic control probe. This new specific binding site was detected by the synthetic control probe (5′-CY5-CTC GCG CAG TGC CTG GAT GAC CCG GG-BHQ1-3′). Genetic analysis For genetic analysis of real-time RT-PCR positive samples, M-segment sequences were amplified using the Superscript III One-Step RT-PCR Kit (Qiagen) according to the manufacturer’s instructions. Primers were M-Sg forw (5′-ATG TATGTTTTATTAACAATT-3′), targeting nt position 21–41 and Gn-rev2 (5′-TTATGCTGATGCATATGAGA
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C-3′) targeting nt position 2073–2090 of the M-segment (according to RVFV strain MP-12; GenBank accession DQ380208.1). The following cycles were used as follows: one cycle at 55°C for 30 min and one cycle at 94°C for 2 min followed by 40 cycles at 94°C for 15 s, 60°C for 30 s and 68°C for 2.5 min followed by a final elongation step at 68°C for 10 min. Following electrophoretic separation, the purified genome fragments were sequenced. Sequence alignment and phylogenetic analysis were performed using Bioedit and MEGA 5 analysis software. RVFV competition ELISA All samples were tested with the ID Screen RVFV competition multispecies ELISA (ID Vet; Montpellier, France) according to the manufacturer’s instructions. Field samples blocking more than 60% of the RVFV detection antibodies compared with a negative control serum (i.e. OD field sample/OD negative control (S/N *100 in %) lower than 40) were qualified ‘positive’. Samples giving results between 40% and 50% were ‘inconclusive’, and results above 50% were considered ‘negative’. All sera were tested in duplicate. RVFV IgM capture ELISA Samples from small ruminants and cattle were also tested with the ID Screen Rift Valley Fever IgM capture ELISA (ID Vet; Montpellier, France) following the instruction manual. Samples producing OD field sample/OD positive control ratios (in%) above 50 were considered to be ‘positive’, those between 40 and 50 ‘inconclusive’ and samples with ratios lower than 40 ‘negative’. Indirect IgG DGn ELISA All samples from sheep and goats were tested in an inhouse indirect IgG ELISA based on bacterially expressed DGn protein. Production of recombinant antigen and the assay procedure was carried out as described previously (J€ackel et al., 2013). Maxisorb immunoplates (Nunc, Roskilde, Denmark) were coated with recombinant RVFV DGn protein, diluted in 0.05 M carbonate–bicarbonate buffer (pH 9.6) to a concentration of 2 lg/ml (100 ll diluted antigen per well). After incubating the plates overnight at 4°C, they were washed three times with 300 ll washing buffer (phosphate-buffered saline (PBS) pH 7.2; 0.1% Tween 20), followed by a blocking step with 200 ll/ well 10% skim milk powder (DIFCOTM) diluted in PBS for 1 h at 37°C in a moist chamber. Polyclonal hyperimmune rabbit serum as positive control was diluted 1 : 20 000 in PBS containing 2% skim milk powder (dilution buffer), and the negative control (negative goat serum) and Mau-
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ritanian serum samples were diluted 1 : 25 in dilution buffer. The positive control was tested in quadruplicate whereas the negative control, conjugate control (2% skim milk) and each sample were tested in duplicate (100 ll per well). Plates were again incubated at 37°C for 1 h in a moist chamber, followed by a further washing step as described above. Horseradish peroxidase-conjugated Protein G (Calbiochem) was diluted 1 : 5000 in dilution buffer, and 100 ll were then added to each well. After another 1 h incubation step, plates were washed again and 100 ll substrate (2,2′-azino di-ethylbenzothiazoline sulphonic acid (ABTS); Roche, Mannheim, Germany) was added to each well. Reactions were run for 30 min at room temperature in the dark, stopped by addition of 1% sodium dodecyl sulphate and the optical density (OD value) at 405 nm was determined. Results were expressed as percentages of the positive control serum (PP value): (mean OD of duplicate test serum/median OD of quadruplicate positive control) 9 100. All samples with PP values lower than the cut-off were considered to be ‘negative’, sera with higher PP values ‘positive’. Indirect immunofluorescence assay All sera were screened for anti-RVFV IgG antibodies by indirect immunofluorescence assay (IIFA) using a modified commercial RVFV IIFA slide test kit (Euroimmun, L€ ubeck, Germany). Slides containing a mixture of infected and non-infected Vero E6 cells on one field (positive field) and non-infected Vero E6 cells on a negative control field were used. Sheep, goat and cattle sera were diluted 1 : 100 with sampling buffer, and 25 ll of the diluted samples were applied to a biochip and incubated for 30 min at room temperature. After a first washing step for 10 min with phosphate-buffered saline (PBS), pH 7.2; 0.2% Tween 20, 25 ll of donkey anti-sheep IgG Cy3 (Indocarbocyanin)-labelled antibodies (Dianova, Hamburg, Germany) for small ruminants and goat anti-bovine IgG Cy3-labelled antibodies (Dianova) diluted 1 : 200 in PBS containing 0.2% Tween were applied. For all small ruminant samples, anti-sheep secondary antibody was used. After a second incubation step at room temperature in the dark for 30 min, the slides were washed again for 10 min, dried and covered with sampling buffer and cover plates. Slides were then examined on a fluorescence microscope (Nikon, Tokyo, Japan). A modification was necessary for testing camel sera as no fluorescence-labelled secondary antibody to camelid IgG was available. Therefore, a polyclonal anti-camel antiserum produced in rabbits (Bethyl laboratories, Montgomery, TX, USA) was used instead (dilution 1 : 100 in sampling buffer) followed by a Cy3 conjugated goat anti-rabbit antibody (Dianova) diluted 1 : 800 in PBS with 0.2% Tween.
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different real-time RT-PCR assays. Specific RVFV RNA could be detected in both assays in four camels (Fig. 2a). All real-time RT-PCR positive samples were collected in the central Mauritanian region of Adrar in the department of Aoujeft. Ct-values ranged from 24.9 to 35.71 (Fig. 2a). The viral load was highest in sample # 8 with 143 270 copies per reaction and Ct-values of 24.9 (assay 1) and 26.22 (assay 2), followed by sample # 9 with more than 7000 copies per reaction and Ct-values of 29.36 (assay 1) and 29.95
Results Quantitative real-time RT-PCR and genetic analysis From December 2010 until February 2011, 163 samples from small ruminants (sheep and goats), cattle and camels were collected in nine different Mauritanian regions throughout the country (Figs 1a and b). The collection included serum samples from 93 small ruminants, 62 camels and eight bovines. All samples were tested with two
(b)
(a)
region
number of sera
small ruminants
camel
cattle
Inchiri/Akjoujt
40
40
-
-
Adrar/Aoujeft
38
28
10
-
Nouadibou
39
-
39
-
Brakna
3
-
3
-
Nouakchott
8
7
-
1
Assaba/Kankossa
1
-
-
1
Gorgol/Mbout
12
2
10
-
Trarza/Ouad Naga
8
2
-
6
Erch
14
14
-
-
Total
163
93
62
8
Fig. 1. (a) Outbreak map of Rift zalley fever virus in Mauritania in 2010. Three regions in northern parts of the country were affected by the virus. (b) Sample origin and numbers. In total, 163 samples were collected from small ruminants (n = 93), camelids (n = 62) and cattle (n = 8) from nine different regions of Mauritania.
(a)
sample
species
region
assay 1
cop/rx
assay 2
sample 8
camel
sample 9
camel
Adrar/Aoujeft
24,9
143270
26,22
Adrar/Aoujeft
29,36
7181
29,95
p 124 sample
camel
Adrar/Aoujeft
34,13
103
35,71
sample 125
camel
Adrar/Aoujeft
34,74
67,9
33,78
(b) 96 DQ380186 OS-1 Mauritania 1987 human 63 54
DQ380184 OS-3 Mauritania 1987 human JQ974833 Mauritania camel NC_014396 Z H548 Egypt 1977 DQ380197 Saudi Arabia 2000 human NC_005220 S23 Uukuniemi
0.1
Bootstrap consensus tree (1000 replicates) Fig. 2. (a) Real-time RT-PCR results from four positive camel samples. Specific viral RNA from Rift Valley fever virus (RVFV) could be detected with two different real-time RT-PCR assays. (b) Phylogenetic analysis of an 859 nucleotide sequence from the M-segment detected in a Mauritanian camel (sample number 8). Sequences were analysed with 1000 replicate bootstrap values using Kimura 2 parameter model. Numbers indicate the percentage of 1000 bootstrap replicates that support each labelled node. For each sequence, the GenBank accession number, strain designation and strain origin are provided. Analysis was supported by the use of Uukuniemi strain S23 as out-group.
34
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(assay 2). The viral load of samples # 124 and # 125 with Ct-values above 33 in both assays was lower with 103 and 68 copies per reaction, respectively. From sample # 8, a partial M-segment sequence of about 859 nucleotides was generated and sequenced [GenBank accession JQ974833; nt position 66–924, according to RVFV strain ZH-548 (GenBank accession NC 014396.1)]. A phylogenetic analysis was performed using sequences from Mauritania 1987, Egypt 1977 and Saudi Arabia 2000. The newly generated camel sequence was closely associated with viruses from the Mauritanian outbreak in 1987 (Fig. 2b). Only three amino acid differences were found between the isolates from 1987 to 2010. However, there were only four different amino acids compared with RVFV strain ZH-548 and seven different amino acids compared with the Saudi Arabian isolate. As all sequences shared a nucleotide homology between 94% and 95%, a more detailed genetic analysis cannot be carried out. Partial sequences derived from the Mauritanian outbreak in 2003 could not be included in the phylogenetic analysis as they overlapped just in a short section (nt 823–924) with the newly generated camel sequence. RVFV competition ELISA and IgM capture ELISA A total of 93 samples (small ruminants n = 64; cattle n = 1; camel n = 28) were positive in the ID Screen RVFV competition multispecies ELISA, which detects IgG as well as IgM antibodies. Four samples gave inconclusive results, and 66 samples were negative (Table 1A; columns 2–4). Overall seroprevalence rates were found in Adrar/Aoujeft (87%), Inchiri/Akjoujt (78%), Nouadibou (41%) and Erch (57%; Table 1B, column 3). The seroprevalence rate among small ruminants was 69% compared with 45% in camels (Table 1A, column 5). Just one bovine sample from eight tested sera was positive in the competition ELISA (Table 1A, column 2). The highest RVF antibody seroprevalence in small ruminants was found in Adrar/Aoujeft (86%) and Inchiri/Akjoujt (78%), whereas affected camels originated mainly from Adrar/Aoujeft and Nouadibou regions (Table 2). A total of 49 samples (all small ruminants) were positive in the ID Screen Rift Valley Fever IgM capture ELISA giving evidence for recent infections by the virus, four samples had a doubtful result and in 48 samples (small ruminants n = 40; cattle n = 8), and no RVFV-specific IgM antibodies were detected (Table 1A, columns 6–8). The highest number of positive small ruminants was found in Inchiri/ Akjoujt with 63%, followed by Adrar/Aoujeft and Erch, both with 57% IgM-positive animals (Table 1B, column 5). In other regions, no further IgM-positive animals were found. All bovine samples were IgM negative (Table 1A, column 9). Camels could not be tested for the presence of
Rift Valley Fever Outbreak in Mauritania
IgM antibodies as the commercial IgM capture ELISA is not functioning in camelids. In-house indirect IgG DGn ELISA All small ruminant samples were also analysed with an in-house indirect IgG DGn-based ELISA. The results are displayed in Table 1A (column 10–12) and Table 1B (columns 6–7). In total, 51 samples were positive with a PP value above 20 752. No RVFV-specific IgG antibodies were detected in the remaining 42 samples. The seroprevalence was 55% with the highest prevalence of IgG antibodies in Adrar/Aoujeft (79%) followed by Inchiri/Akjoujt (60%), Erch (29%) and Nouakchott (14%). There were no IgGpositive animals in Trarza and Gorgol. Comparison of all three ELISA assays revealed additional results for the infection status of small ruminants (Table 3): 36 of the 51 DGn ELISA-positive samples were also positive with both commercial ELISAs. Eleven samples showed evidence of a past infection as they were just positive with the IgG DGn-based ELISA and with the ID Screen competition ELISA. In 13 animals, a recent infection could be shown as they were just positive with the two commercial ELISAs, both detecting IgM antibodies. Indirect immunofluorescence assay All samples were eventually tested in an IIFA for the presence of RVFV-specific IgG antibodies (using speciesspecific secondary antibodies). As shown in Fig. 3, in sera of all four species, RVF virus antibodies were detected [see Figs 3a (sheep), b (goat), c (camel) and d (cattle)]. With regard to small ruminants, 57 sera showed positive staining; eight had a doubtful result and no RVFV-specific IgG antibodies were detected in 28 samples (Table 1A; columns 13–16). Just two of eight cattle samples were positive in indirect immunofluorescence; in the remaining six samples, no IgG antibodies could be detected. 27 camels were positive, 11 sera had a doubtful result, whereas in 24 samples no antibodies were found (Table 1A; columns 13–16). Prevalence was highest in Adrar/Aoujeft with 82% (Table 1B, columns 8–9). In Inchiri/Akjoujt, the seroprevalence was 75%, for Erch it was lower (43%). In Gorgol/Mbout and Brakna, the seroprevalence in the immunofluorescence assay was 33%, followed by Nouadibou with 31% and Trarza/Ouad Naga where two of eight tested animals were positive (25%). In Nouakchott and Assaba/Kankossa, no IIFApositive animals were found (Table 1B, columns 8–9). Discussion In October/November 2010, a massive outbreak of Rift Valley fever occurred in northern Mauritania after unusually
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35
36 32 7 66
27
Negative
45 13 57
69
Prevalence (%)
31/40 33/38 16/39 1/3 1/8 0/1 2/12 1/8 8/14 93/163
Inchiri/Akjoujt Adrar/Aoujeft Nouadibou Brakna Nouakchott Assaba/Kankossa Gorgol/Mbout Trarza/Ouad Naga Erch Total
78 87 41 33 13 0 17 13 57 57
Prevalence (%)
n.t. 0 49
49 n.t. 0 4
4
Doubtful
n.t. 8 48
40
Negative
n.t. n.t. 51
– 0 59
25/40 16/28 n.t. n.t. 0/8 0/1 0/2 0/8 8/14 49/101
Positive/total 63 57 n.t. n.t. 0 0 0 0 57 49
Prevalence (%)
n.t. n.t. 42
42
Negative
– – 55
55
Prevalence (%)
24/40 22/28 n.t. n.t. 1/7 n.t. 0/2 0/2 4/14 51/93
Positive/total
27 2 86
57
Positive
60 79 n.t. n.t. 14 n.t. 0 0 29 55
24 6 58
28
Negative
a
44 25 53
61
Prevalence (%)
30/40 31/38 12/39 1/3 0/8 0/1 4/12 2/8 6/14 86/163
Positive/total
75 82 31 33 0 0 33 25 43 53
Prevalence (%)
Indirect immunofluorescence assay
11 0 19
8
Doubtful
Indirect immunofluorescence assay
Prevalence (%)
Indirect IgG Gn ELISAb
51
Positive
Indirect IgG Gn ELISAb
53
Prevalence (%)
ID screen IgM capture ELISAa
Positive
ID screen IgM capture ELISAa
The number of positive, negative and doubtful samples for each assay and species as well as the resulting seroprevalence is shown. The percentages were rounded off to whole numbers. Camels not tested. b Camels and cattle not tested.
Positive/total
Region
ID screen competition ELISA
2 0 4
28 1 93
(B)
2
64
Small ruminants Camel Cattle Total
Doubtful
Positive
ID screen competition ELISA
Species
(A)
Table 1. Serological analysis of Mauritanian serum samples from small ruminants, cattle and camels with three different ELISAs (ID Vet ID Screen Rift Valley Fever competition ELISA (multispecies); ID Vet ID Screen Rift Valley Fever IgM capture ELISA and in-house indirect IgG DGn ELISA) and indirect immunofluorescence assay (IIFA). Camels were only tested with the ID VET competition ELISA and IIFA, cattle samples were not tested with the in-house indirect IgG ELISA. (A) Results of the four assays grouped into species. (B) Results of the four assays with regard to geographic distribution
Rift Valley Fever Outbreak in Mauritania S. J€ackel et al.
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Rift Valley Fever Outbreak in Mauritania
Table 2. Seroprevalence of IgG/IgM antibodies in different Mauritanian regions determined with the ID VET competition ELISA subdivided into species. Small ruminants
Cattle
Region
Positive/ total
%
Inchiri/Akjoujt Adrar/Aoujeft Nouadibou Brakna Nouakchott Assaba/Kankossa Gorgol/Mbout Trarza/Ouad Naga Erch Total
31/40 24/28 – – 1/7 – 0/2 0/2 8/14 64/93
78 86 – – 14 – 0 0 57 69
Camels
Positive/ total
%
– – – – 0/1 0/1 – 1/6
– – – – 0 0 – 17
1/8
13
Positive/ total
%
– 9/10 16/39 1/3 – – 2/10 – – 28/62
– 90 41 33 – – 20 – – 45
long and heavy rainfalls. Humans, livestock and camels were affected (El Mamy et al., 2011). In the here presented follow-up study, we assayed field samples collected in the late time period of this epidemic (December 2010 to February 2011) to monitor whether RVFV still circulated in animals at this time. As all samples in this as well as in an earlier study (El Mamy et al., 2011) were regular submissions to the diagnostic laboratory, they do not represent a random selection, that is, results need to be interpreted carefully. In our study, sera from livestock were tested for viremia by real-time qRT-PCR and run in two commercially available ELISAs. Moreover, small ruminant sera were also tested in a newly developed indirect IgG ELISA (J€ackel et al., 2013). Our results show that the overall seroprevalence rate for IgG/IgM antibodies (as measured by ID VET competition ELISA) in livestock animals and camels increased from 37% to 57% when compared with recently published
Table 3. Identification of the serological status of small ruminants by comparing three different ELISAs. Samples with a doubtful result in competition ELISA and/or IgM capture ELISA are not shown
ELISA result
Competition ELISA positive IgM capture ELISA postive Indirect IgG Gn ELISA positive
Competition ELISA positive IgM capture ELISA postive Indirect IgG Gn ELISA negative
Competition ELISA positive IgM capture ELISA negative Indirect IgG Gn ELISA positive
Competition ELISA positive IgM capture ELISA negative Indirect IgG Gn ELISA negative
Sample number
36 (IgM and IgG)
13 (IgM only)
11 (IgG only)
27 (negative)
(a)
(b)
Mau123 (c)
Mau10 (d)
Mau7
Mau146
€beck, Germany) Fig. 3. Indirect immunofluorescence assay (IIFA) using a commercial Rift Valley fever virus (RVFV) IIFA slide test kit (Euroimmun, Lu with slight modifications for testing ruminant and camel serum samples. (a) sheep, (b) goat, (c) camel, (d) cattle.
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S. J€ackel et al.
Rift Valley Fever Outbreak in Mauritania
results generated from sera sampled in September and October 2010 (El Mamy et al., 2011). In particular, the seroprevalence in small ruminants rose from 43% to 69% over this time. Furthermore, there was an increase in the number of IgM-positive animals (53% compared with 42%). Used in combination, the three assays demonstrated that only 11 of 93 small ruminants showed exclusively IgG antibodies, which points to a recent virus exposure of the majority of sampled animals. The IIFA, which was originally designed for testing human sera, was adapted to animal sera and yielded results, which clearly correlated with the ELISA results. Camels played an important role in RVFV epidemics in Kenya (Scott et al., 1963; Davies et al., 1985), Nigeria (Davies et al., 1985), Sudan (Eisa, 1984), Egypt (Davies et al., 1985; Ahmed Kamal, 2011) and Morocco (El-Harrak et al., 2011). During the first RVF outbreak in Egypt, camels which were moved from northern Sudan to the southern parts of Egypt may have spread the virus (Ahmed Kamal, 2011). In Morocco, El-Harrak et al. found a seroprevalence of 15% among camels in 2009 and suspected animal movements from south-eastern parts of the Sahara desert to the north-west as reasons. After an earlier outbreak in Mauritania in 1987, camels carried RVF antibodies at a rate of about 33% (Saluzzo et al., 1987). In 1998, Nabeth et al. found one IgM-positive sample of 39 tested camels. Camels also played a very important role in the RVF outbreak in Mauritania in 2010, and numerous animals showed clinical signs. Rift Valley fever virus RNA was detected in seven animals by real-time RT-PCR, and virus was isolated from four of these samples (El Mamy et al., 2011). The overall seroprevalence rate for IgG/IgM antibodies in camels was again about 33%. In the here described study on late epidemic sera, a seroprevalence rate of about 45% was found in camels, which underlines their central role during the latest Mauritanian outbreak. Moreover, using two different quantitative real-time RT-PCR assays, we detected RVFV RNA in four camelid samples and a partial sequence of the M-segment was obtained from one animal (GenBank accession number: JQ974833). To our knowledge, this is the first published camel-derived RVFV sequence apart from sequences originating from ticks feeding on a camel in 1979 (accession numbers: HQ403564.1; HQ403562.1; HQ403563.1). The phylogenetic analysis showed that this sequence is closely associated with those of viruses from the Mauritanian outbreak in 1987. However, due to the low variability and a high nucleotide homology of >95% to other virus sequences, further genetic analysis of this fragment was not possible. In conclusion, we could show that the RVF virus was circulating in Mauritania after the initial outbreak in October 2010 at least until January 2011, when the epidemic was officially declared to be over, since two camels were still viremic 38
then. Furthermore, our results highlight the important role of small ruminants during the infection cycle of RVFV. Acknowledgements We would like to acknowledge Dr. Modou Moustapha L^ o, L’Institut senegalais de recherches agricoles (ISRA) / Laboratoire National de l’Elevage et de Recherches Veterinaires (LNERV), Dakar, Senegal, Dr. Anne Balkema-Buschmann, Friedrich-Loeffler-Institut and Dr. Hermann Unger, Joint FAO/IAEA Division, Vienna, Austria, for their scientific and technical support of this study. This study was partially funded by EU grant FP7-261504 EDENext and is catalogued by the EDENext Steering Committee as EDENext159 (\http://www.edenext.eu) as well as by the FP7 NADIR project. The contents of this publication are the sole responsibility of the authors and do not necessarily reflect the views of the European Commission. Conflicts of interest The authors have no conflicts of interest to declare. References Ahmed Kamal, S., 2011: Observations on rift valley fever virus and vaccines in Egypt. Virol. J. 8, 532. Bird, B. H., D. A. Bawiec, T. G. Ksiazek, T. R. Shoemaker, and S. T. Nichol, 2007: Highly sensitive and broadly reactive quantitative reverse transcription-PCR assay for high-throughput detection of Rift Valley fever virus. J. Clin. Microbiol. 45, 3506–3513. Bird, B. H., T. G. Ksiazek, S. T. Nichol, and N. J. MacLachlan, 2009: Rift Valley fever virus. J. Am. Vet. Med. Assoc. 234, 883–893. Bouloy, M., and F. Weber, 2010: Molecular biology of rift valley fever virus. Open Virol. J. 4, 8–14. Chevalier, V., T. Rakotondrafara, M. Jourdan, J. M. Heraud, H. R. Andriamanivo, B. Durand, J. Ravaomanana, P. E. Rollin, and R. Rakotondravao, 2011: An unexpected recurrent transmission of Rift Valley fever virus in cattle in a temperate and mountainous area of Madagascar. PLoS Negl. Trop. Dis. 5, e1423. Daubney, R., J. R. Hudson, and P. C. Garnham, 1931: Enzootic hepatitis or rift valley fever. An undescribed virus disease of sheep, cattle and man from East Africa. J. Pathol. Bacteriol. 34, 545–579. Davies, F. G., J. Koros, and H. Mbugua, 1985: Rift Valley fever in Kenya: the presence of antibody to the virus in camels (Camelus dromedarius). J. Hyg. (Lond) 94, 241–244. Digoutte, J. P., and C. J. Peters, 1989: General aspects of the 1987 Rift Valley fever epidemic in Mauritania. Res. Virol. 140, 27–30. Eisa, M., 1984: Preliminary survey of domestic animals of the Sudan for precipitating antibodies to Rift Valley fever virus. J. Hyg. (Lond) 93, 629–637.
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