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Early detection of viral excretion from experimentally infected goats with peste-des-petits ruminants virus
Preventive Veterinary Medicine 78 (2007) 85–88 www.elsevier.com/locate/prevetmed
Short communication
Early detection of viral excretion from experimentally infected goats with peste-des-petits ruminants virus E. Couacy-Hymann a,*, S.C. Bodjo a,c, T. Danho a, M.Y. Koffi a, G. Libeau b, A. Diallo c a
LANADA/Laboratoire Central de Pathologie Animale de Bingerville, BP: 206 Bingerville, Cote-d’Ivoire b CIRAD/EMVT, Campus International de Baillarguet, 34398 Montpellier, Cedex 5, France c Animal Production Unit, FAO/IAEA Agriculture and Biotechnology Laboratory, Laboratories, A-2444 Seibersdorf, Austria Received 15 January 2006; received in revised form 11 July 2006; accepted 20 September 2006
Abstract We observed 15 goats for 9 days after subcutaneous infection with 103 TCID50 with isolates of peste-des-petits ruminants virus from Africa and India and five concurrent, uninfected control goats. Typical clinical signs of the infection were present in all 15 infected goats by day 8 and in most by day 6 and some signs were present by day 4. However, 6 out of 15 goats already have detectable virus shedding by day 3 and four more were shedding by day 4 and every goat had virus shedding for at least 1 day before the recognition of clinical signs. This experiment indicates that incubatory carriers therefore might play a role in the transmission of PPRV among small ruminants. # 2006 Elsevier B.V. All rights reserved. Keywords: Peste-des-petits-ruminants (PPR); PPRV; Morbillivirus; Goat; Epidemiology
1. Introduction Peste-des-petits-ruminants (PPR) is a disease of sheep and goats with high morbidity and mortality; it is one of the major notifiable diseases of the World Organisation for * Corresponding author. Tel.: +225 22 403 136/138; fax: +225 22 403 644. E-mail address: [email protected] (E. Couacy-Hymann). 0167-5877/$ – see front matter # 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.prevetmed.2006.09.003
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E. Couacy-Hymann et al. / Preventive Veterinary Medicine 78 (2007) 85–88
Animal Health (OIE). It has a widespread distribution across Sub-Saharan Africa, the Middle-East and Southern Asia, where it still causes serious economic losses (Taylor, 1984a; Shaila et al., 1989; Diallo, 2003) and remains a major constraint on the development of small-ruminant farms in these countries. Outbreaks of PPR are recorded regularly by field technicians and farmers (mainly during the raining season) in tropical regions and are characterized by fever, erosive stomatitis, nasal and ocular discharges, pneumonia that could be exacerbated by secondary bacterial infection and diarrhoae. The severity of the clinical signs depends on the virulence of the strains (Couacy-Hymann, unpublished data). The causative agent is peste-des-petits-ruminants virus (PPRV), a member of the Morbillivirus genus in the Paramyxoviridae family (Gibbs et al., 1979). PPRV is closely related to rinderpest virus (RPV) which infects cattle and other large ruminants and can cause disease in small ruminants (Anderson et al., 1990; Couacy-Hymann et al., 1995; Diallo, 2003). PPRV is transmitted from infected animals to naı¨ve ones by close contact because it is readily inactivated in external conditions. The oral and respiratory tracts are the main routes of virus tranmission through oral, nasal and ocular excretions and the virus is rapidly inactivated in dead animals. There is no chronic or convalescent carrier form of this infection. However, rinderpest (RP) virus (RPV) is shed by incubatory carriers (Plowright, 1968; Scott et al., 1986). Such carriers contribute to the transmission of RPV. Because RPV and PPRV are closely related, our objective was to investigate whether PPRV was shed prior to clinical signs of PPR in infected goats. For this purpose, 20 West African dwarf goats, aged between 2 and 3 years and weighting between 12 and 15 kg, were purchased from villages located in the rain forest region and tested as negative for the presence of antibodies against RP and PPR by the virus-neutralization test (Rossiter and Jessett, 1982; Taylor, 1984b). Each group was housed in its own cage with its own feeding and drinking tanks. Each animal was treated (300 mg/kg) with Albendazole, an anti-parasite drug, during the acclimatisation period.Five virus isolates were used for the challenge. They were obtained from the virus bank of CIRAD/EMVT, Montpellier, France and represented different geographical regions within Africa and India: Coˆte-d’Ivoire 89 (CI89); Conakry Guinea (CG); Nigeria 75/1 (NIG 75/1); Sudan-Sennar (SS); India-Calcutta (IC). We used a table of random numbers to divide the 20 goats into six groups: five groups each of three goats and one control group of five goats. Each animal (except controls) was infected subcutaneously with 1 ml of the various challenge viral suspensions, at a concentration of 103 TCID50 ml 1. Each group of animals were kept separately in an 8-m2 specific-cage, in the laboratory-animal house. Each cage was identified with a specific number and had a separate attendant blindly assigned to feed and water the animals. Animals were examined daily and rectal temperature was daily recorded. All infected and control animals were slaughtered at 9 day. The first author was certificated in 2004 by the International Committee for Laboratory Animal Sciences (ICLAS). Nasal, ocular (conjunctival) fluids and saliva were collected daily using cotton swabs from all challenged animals from day 1 to day 9 post-infection. Samples also were collected from the controls on day 5 and 9. All samples were stored at 80 8C until examined. These samples were processed blinded to the treatment group, at the end of the experiment, for RNA extraction using the Qiagen RNA Extraction kit (Qiagen, RNEasy
E. Couacy-Hymann et al. / Preventive Veterinary Medicine 78 (2007) 85–88
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Table 1 Days post-infection on which clinical signs and positive RT-PCR tests for peste-des-petits-ruminants (PPR) were present in 15 experimentally infected goats Strain and animal
Temperature Nasal/ocular Oral Diarrhoea RT-PCR Results 39 8C discharges ulceration 1-2 3 4
5
6–9
Coˆte a 5–8 D’Ivoire 89 b 6–7 c 5–8
4–9 6–8 4–8
7 5 –
7–9 6–8 6–8
– – –
+O/N +O/N/S +O/N/S +O/N/S – +O/N/S +O/N/S +O/N/S +O/N +O/N/S +O/N/S +O/N/S
Guinea Conakry
a 7–8 b 6–8 c 6–7
7 7–9 8–9
5 7 –
– 7–9 6–8
– – –
+O – +O
+O/N/S +O/N/S +O/N/S +O/N/S +O/N/S +O/N/S +O/N/S +O/N +O/N/S
Nigeria 75/1
a 6–8 b 7–8 c 8
– 8 –
– – –
9 – –
– – –
– – –
– – –
+O/N +O –
+O/N/S +O/N/S +O/N/S
Sudan-Sennar a 7–8 b 6–7 c 8
6 7–8 –
– 8 –
– 8–9 –
– –
– – –
– +O –
+O +O/S –
+O/N/S +O/N/S +O/N/S
India-Calcutta a 8 b 7–9 c 6–8
– 7–9 9
– 7 8
– 8–9 –
– – –
– +O +O +O/N/S +O/N +O/N/S +O/N/S +O/N/S +O/N +O/N +O/N/S +O/N/S
(–), No clinical sign/negative RT-PCR results; +O/N/S, positive result on ocular, nasal and/or saliva samples.
Mini kit, USA) following the manufacturer’s recommended protocol. Within each group, each day for each type of sample, 20 ml of RNA solution from the three goats were pooled. Then they were analysed using the RT-PCR technique according to the methodology outlined by Couacy-Hymann (1994) and Couacy-Hymann et al. (2002) to amplify a DNA fragment of 296 bp targeting the nucleoprotein (Np) gene. When a pool was positive, its individual samples (RNA solution) were used to find out which goats were positive. The clinical responses of the goats to the infections and the RT-PCR results are in Table 1. All 15 infected animals (and none of the five controls) showed clinical signs of PPR. Some goats had clinical signs as early as day 4, and only two goats did not have signs by day 6. However, we detected virus at least in the ocular samples as early as day 3 in 6 out of 15 goats, and in every goat at least 1 day (and up to 4 days) before the goat showed its earliest clinical sign. None of the control goats ever had a positive RT-PCR result demonstrating that there was no natural circulation of PPRV in the laboratory-animal house environment. After a transfer of the target amplified fragments onto a membrane positively charged by southern-blotting, the probe SP1, internal to the target sequence, confirmed all positive and negative results obtained with RT-PCR, from the samples collected from the infected goats from days 0 to 5. For the samples collected from those goats from days 6 to 9, we did not analyse the individual-goat samples because of the strong evidence that every goat was shedding; however, all pooled samples were positive across those days. Here, too, samples from control goats remained negative. Although we did not do any infectivity studies, the RT-PCR evidence carrier state suggests that there is an incubatory-carrier state in goats infected with PPRV. Our clinical
88
E. Couacy-Hymann et al. / Preventive Veterinary Medicine 78 (2007) 85–88
signs data are in accord with our previous (unpublished) studies of the virulence of these strains. We used five different isloates to check whether the virulent strains (Coˆte-d’Ivoire 89, Guinea Conakry) spread infectious particles earlier than the mild ones (Nigeria 75/1, Sudan-Sennar). The Nigeria 75/1 strain is used as a homologous PPR vaccine strain (Diallo et al., 1989). The RT-PCR technique used has been shown to be more sensitive than using virus isolation and specific to detect Np gene of PPRV (Couacy-Hymann et al., 2002). Whether incubatory carriers of PPRV indeed exist, and small ruminants can transmit the infection before clinical suspicion of the disease would be present, of course needs to be confirmed.
Acknowledgements I gratefully thank Dr. John Crowther (Joint FAO/IAEA) for his support. This study has been funded by a grant from EU (Contract, ICA4 CT-2000 30027).
References Anderson, E.C., Hassan, A., Barrett, T., Anderson, J., 1990. Observations on the pathogenicity for sheep and goats and the transmissibility of the strain of virus isolated during the rinderpest outbreak in Sri-Lanka in 1987. Vet. Microbiol. 21, 309–318. Couacy-Hymann, E., Roger, F., Huard, C., Guillou, J.P., Libeau, G., Diallo, A., 2002. Rapid and sensitive detection of peste des petits ruminants virus by a polymerase chain reaction assay. J. Virol. Methods 100, 17–25. Couacy-Hymann, E., Bidjeh, K., Angba, A., Domenech, J., Diallo, A., 1995. Protection of goats against rinderpest by vaccination with attenuated peste des petits ruminants virus. Res. Vet. Sci. 59, 106–109. Couacy-Hymann, E. 1994. La lutte contre la peste bovine en Coˆte-d’Ivoire. Couˆts et Be´ne´fices des Campagnes de Prophylaxie. Proble`mes pose´s pour son Eradication. The`se de Doctorat d’universite´. Universite´ Paris XII-Val de Marne-Cre´teil, vols. I et II. Diallo, A., 2003. Control of PPR: classical and new generation of vaccine. Dev. Biol. Basel, Karger 114, 85–91. Diallo, A., Taylor, W.P., Lefevre, P.C., Provost, A., 1989. Atte´nuation d’une souche de virus de la peste des petits ruminants: candidat pour un vaccin homologue vivant. Revue d’Elevage et de Me´decine Ve´te´rinaire des Pays Tropicaux 42, 311–319. Gibbs, E.P.J., Taylor, W.P., Lawman, M.P.J., Bryant, J., 1979. Classification of peste des petits ruminants virus as a fourth member of the genus Morbillivirus. Intervirology 11, 268–274. Plowright, W., 1968. Rinderpest virus. Virology Monographs, vol. 3. Spring-Verlag, Wien, New York, pp. 25–110. Rossiter, P.B., Jessett, D.M., 1982. Microtitre technique for the assay of rinderpest virus and neutralising antibody. Res. Vet. Sci. 32, 253–256. Scott, G.R., Taylor, W.P., Rossiter, P.B. 1986. Manual of the diagnosis of rinderpest. FAO Animal Health Manual, first ed. Rome, 215 p. Shaila, M.S., Purushothaman, V., Bhavasar, D., Venugopal, K., Venkatesan, R.A., 1989. Peste des petits ruminants in India. Vet. Rec. 125, 602. Taylor, W.P., 1984a. The distribution and epidemiology of peste des petits ruminants. Prev. Vet. Med. 2, 157–166. Taylor, W.P., 1984b. A microneutralisation test for detection of rinderpest antibody. Revued’Elevage et de Me´decine Ve´te´rinaire des pays Tropicaux 37, 155–159.
Short communication
Early detection of viral excretion from experimentally infected goats with peste-des-petits ruminants virus E. Couacy-Hymann a,*, S.C. Bodjo a,c, T. Danho a, M.Y. Koffi a, G. Libeau b, A. Diallo c a
LANADA/Laboratoire Central de Pathologie Animale de Bingerville, BP: 206 Bingerville, Cote-d’Ivoire b CIRAD/EMVT, Campus International de Baillarguet, 34398 Montpellier, Cedex 5, France c Animal Production Unit, FAO/IAEA Agriculture and Biotechnology Laboratory, Laboratories, A-2444 Seibersdorf, Austria Received 15 January 2006; received in revised form 11 July 2006; accepted 20 September 2006
Abstract We observed 15 goats for 9 days after subcutaneous infection with 103 TCID50 with isolates of peste-des-petits ruminants virus from Africa and India and five concurrent, uninfected control goats. Typical clinical signs of the infection were present in all 15 infected goats by day 8 and in most by day 6 and some signs were present by day 4. However, 6 out of 15 goats already have detectable virus shedding by day 3 and four more were shedding by day 4 and every goat had virus shedding for at least 1 day before the recognition of clinical signs. This experiment indicates that incubatory carriers therefore might play a role in the transmission of PPRV among small ruminants. # 2006 Elsevier B.V. All rights reserved. Keywords: Peste-des-petits-ruminants (PPR); PPRV; Morbillivirus; Goat; Epidemiology
1. Introduction Peste-des-petits-ruminants (PPR) is a disease of sheep and goats with high morbidity and mortality; it is one of the major notifiable diseases of the World Organisation for * Corresponding author. Tel.: +225 22 403 136/138; fax: +225 22 403 644. E-mail address: [email protected] (E. Couacy-Hymann). 0167-5877/$ – see front matter # 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.prevetmed.2006.09.003
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E. Couacy-Hymann et al. / Preventive Veterinary Medicine 78 (2007) 85–88
Animal Health (OIE). It has a widespread distribution across Sub-Saharan Africa, the Middle-East and Southern Asia, where it still causes serious economic losses (Taylor, 1984a; Shaila et al., 1989; Diallo, 2003) and remains a major constraint on the development of small-ruminant farms in these countries. Outbreaks of PPR are recorded regularly by field technicians and farmers (mainly during the raining season) in tropical regions and are characterized by fever, erosive stomatitis, nasal and ocular discharges, pneumonia that could be exacerbated by secondary bacterial infection and diarrhoae. The severity of the clinical signs depends on the virulence of the strains (Couacy-Hymann, unpublished data). The causative agent is peste-des-petits-ruminants virus (PPRV), a member of the Morbillivirus genus in the Paramyxoviridae family (Gibbs et al., 1979). PPRV is closely related to rinderpest virus (RPV) which infects cattle and other large ruminants and can cause disease in small ruminants (Anderson et al., 1990; Couacy-Hymann et al., 1995; Diallo, 2003). PPRV is transmitted from infected animals to naı¨ve ones by close contact because it is readily inactivated in external conditions. The oral and respiratory tracts are the main routes of virus tranmission through oral, nasal and ocular excretions and the virus is rapidly inactivated in dead animals. There is no chronic or convalescent carrier form of this infection. However, rinderpest (RP) virus (RPV) is shed by incubatory carriers (Plowright, 1968; Scott et al., 1986). Such carriers contribute to the transmission of RPV. Because RPV and PPRV are closely related, our objective was to investigate whether PPRV was shed prior to clinical signs of PPR in infected goats. For this purpose, 20 West African dwarf goats, aged between 2 and 3 years and weighting between 12 and 15 kg, were purchased from villages located in the rain forest region and tested as negative for the presence of antibodies against RP and PPR by the virus-neutralization test (Rossiter and Jessett, 1982; Taylor, 1984b). Each group was housed in its own cage with its own feeding and drinking tanks. Each animal was treated (300 mg/kg) with Albendazole, an anti-parasite drug, during the acclimatisation period.Five virus isolates were used for the challenge. They were obtained from the virus bank of CIRAD/EMVT, Montpellier, France and represented different geographical regions within Africa and India: Coˆte-d’Ivoire 89 (CI89); Conakry Guinea (CG); Nigeria 75/1 (NIG 75/1); Sudan-Sennar (SS); India-Calcutta (IC). We used a table of random numbers to divide the 20 goats into six groups: five groups each of three goats and one control group of five goats. Each animal (except controls) was infected subcutaneously with 1 ml of the various challenge viral suspensions, at a concentration of 103 TCID50 ml 1. Each group of animals were kept separately in an 8-m2 specific-cage, in the laboratory-animal house. Each cage was identified with a specific number and had a separate attendant blindly assigned to feed and water the animals. Animals were examined daily and rectal temperature was daily recorded. All infected and control animals were slaughtered at 9 day. The first author was certificated in 2004 by the International Committee for Laboratory Animal Sciences (ICLAS). Nasal, ocular (conjunctival) fluids and saliva were collected daily using cotton swabs from all challenged animals from day 1 to day 9 post-infection. Samples also were collected from the controls on day 5 and 9. All samples were stored at 80 8C until examined. These samples were processed blinded to the treatment group, at the end of the experiment, for RNA extraction using the Qiagen RNA Extraction kit (Qiagen, RNEasy
E. Couacy-Hymann et al. / Preventive Veterinary Medicine 78 (2007) 85–88
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Table 1 Days post-infection on which clinical signs and positive RT-PCR tests for peste-des-petits-ruminants (PPR) were present in 15 experimentally infected goats Strain and animal
Temperature Nasal/ocular Oral Diarrhoea RT-PCR Results 39 8C discharges ulceration 1-2 3 4
5
6–9
Coˆte a 5–8 D’Ivoire 89 b 6–7 c 5–8
4–9 6–8 4–8
7 5 –
7–9 6–8 6–8
– – –
+O/N +O/N/S +O/N/S +O/N/S – +O/N/S +O/N/S +O/N/S +O/N +O/N/S +O/N/S +O/N/S
Guinea Conakry
a 7–8 b 6–8 c 6–7
7 7–9 8–9
5 7 –
– 7–9 6–8
– – –
+O – +O
+O/N/S +O/N/S +O/N/S +O/N/S +O/N/S +O/N/S +O/N/S +O/N +O/N/S
Nigeria 75/1
a 6–8 b 7–8 c 8
– 8 –
– – –
9 – –
– – –
– – –
– – –
+O/N +O –
+O/N/S +O/N/S +O/N/S
Sudan-Sennar a 7–8 b 6–7 c 8
6 7–8 –
– 8 –
– 8–9 –
– –
– – –
– +O –
+O +O/S –
+O/N/S +O/N/S +O/N/S
India-Calcutta a 8 b 7–9 c 6–8
– 7–9 9
– 7 8
– 8–9 –
– – –
– +O +O +O/N/S +O/N +O/N/S +O/N/S +O/N/S +O/N +O/N +O/N/S +O/N/S
(–), No clinical sign/negative RT-PCR results; +O/N/S, positive result on ocular, nasal and/or saliva samples.
Mini kit, USA) following the manufacturer’s recommended protocol. Within each group, each day for each type of sample, 20 ml of RNA solution from the three goats were pooled. Then they were analysed using the RT-PCR technique according to the methodology outlined by Couacy-Hymann (1994) and Couacy-Hymann et al. (2002) to amplify a DNA fragment of 296 bp targeting the nucleoprotein (Np) gene. When a pool was positive, its individual samples (RNA solution) were used to find out which goats were positive. The clinical responses of the goats to the infections and the RT-PCR results are in Table 1. All 15 infected animals (and none of the five controls) showed clinical signs of PPR. Some goats had clinical signs as early as day 4, and only two goats did not have signs by day 6. However, we detected virus at least in the ocular samples as early as day 3 in 6 out of 15 goats, and in every goat at least 1 day (and up to 4 days) before the goat showed its earliest clinical sign. None of the control goats ever had a positive RT-PCR result demonstrating that there was no natural circulation of PPRV in the laboratory-animal house environment. After a transfer of the target amplified fragments onto a membrane positively charged by southern-blotting, the probe SP1, internal to the target sequence, confirmed all positive and negative results obtained with RT-PCR, from the samples collected from the infected goats from days 0 to 5. For the samples collected from those goats from days 6 to 9, we did not analyse the individual-goat samples because of the strong evidence that every goat was shedding; however, all pooled samples were positive across those days. Here, too, samples from control goats remained negative. Although we did not do any infectivity studies, the RT-PCR evidence carrier state suggests that there is an incubatory-carrier state in goats infected with PPRV. Our clinical
88
E. Couacy-Hymann et al. / Preventive Veterinary Medicine 78 (2007) 85–88
signs data are in accord with our previous (unpublished) studies of the virulence of these strains. We used five different isloates to check whether the virulent strains (Coˆte-d’Ivoire 89, Guinea Conakry) spread infectious particles earlier than the mild ones (Nigeria 75/1, Sudan-Sennar). The Nigeria 75/1 strain is used as a homologous PPR vaccine strain (Diallo et al., 1989). The RT-PCR technique used has been shown to be more sensitive than using virus isolation and specific to detect Np gene of PPRV (Couacy-Hymann et al., 2002). Whether incubatory carriers of PPRV indeed exist, and small ruminants can transmit the infection before clinical suspicion of the disease would be present, of course needs to be confirmed.
Acknowledgements I gratefully thank Dr. John Crowther (Joint FAO/IAEA) for his support. This study has been funded by a grant from EU (Contract, ICA4 CT-2000 30027).
References Anderson, E.C., Hassan, A., Barrett, T., Anderson, J., 1990. Observations on the pathogenicity for sheep and goats and the transmissibility of the strain of virus isolated during the rinderpest outbreak in Sri-Lanka in 1987. Vet. Microbiol. 21, 309–318. Couacy-Hymann, E., Roger, F., Huard, C., Guillou, J.P., Libeau, G., Diallo, A., 2002. Rapid and sensitive detection of peste des petits ruminants virus by a polymerase chain reaction assay. J. Virol. Methods 100, 17–25. Couacy-Hymann, E., Bidjeh, K., Angba, A., Domenech, J., Diallo, A., 1995. Protection of goats against rinderpest by vaccination with attenuated peste des petits ruminants virus. Res. Vet. Sci. 59, 106–109. Couacy-Hymann, E. 1994. La lutte contre la peste bovine en Coˆte-d’Ivoire. Couˆts et Be´ne´fices des Campagnes de Prophylaxie. Proble`mes pose´s pour son Eradication. The`se de Doctorat d’universite´. Universite´ Paris XII-Val de Marne-Cre´teil, vols. I et II. Diallo, A., 2003. Control of PPR: classical and new generation of vaccine. Dev. Biol. Basel, Karger 114, 85–91. Diallo, A., Taylor, W.P., Lefevre, P.C., Provost, A., 1989. Atte´nuation d’une souche de virus de la peste des petits ruminants: candidat pour un vaccin homologue vivant. Revue d’Elevage et de Me´decine Ve´te´rinaire des Pays Tropicaux 42, 311–319. Gibbs, E.P.J., Taylor, W.P., Lawman, M.P.J., Bryant, J., 1979. Classification of peste des petits ruminants virus as a fourth member of the genus Morbillivirus. Intervirology 11, 268–274. Plowright, W., 1968. Rinderpest virus. Virology Monographs, vol. 3. Spring-Verlag, Wien, New York, pp. 25–110. Rossiter, P.B., Jessett, D.M., 1982. Microtitre technique for the assay of rinderpest virus and neutralising antibody. Res. Vet. Sci. 32, 253–256. Scott, G.R., Taylor, W.P., Rossiter, P.B. 1986. Manual of the diagnosis of rinderpest. FAO Animal Health Manual, first ed. Rome, 215 p. Shaila, M.S., Purushothaman, V., Bhavasar, D., Venugopal, K., Venkatesan, R.A., 1989. Peste des petits ruminants in India. Vet. Rec. 125, 602. Taylor, W.P., 1984a. The distribution and epidemiology of peste des petits ruminants. Prev. Vet. Med. 2, 157–166. Taylor, W.P., 1984b. A microneutralisation test for detection of rinderpest antibody. Revued’Elevage et de Me´decine Ve´te´rinaire des pays Tropicaux 37, 155–159.