5 Peste des Petits Ruminants Virus The Pirbright Institute, Compton Laboratory, Compton, UK Peste des petits ruminants (PPR) is a highly contagious, fatal and economically important disease of both domestic and wild small ruminants, and camels. Owing to high morbidity (100%) and mortality (90%), PPR was included in the OIE (Office International des Epizooties) list of notifiable terrestrial animal diseases. The disease is currently spreading rapidly in most countries of the sub-Saharan and North Africa, the Middle East and Indian sub-continent and as far as into Tibet, China. During the rinderpest virus (RPV) eradication campaign, there have been significant improvements in understanding the biology of viruses; however, the primary focus has remained developing and improving efficient vaccines. The present chapter aims to provide an overview of all known features of the PPRV genome, structure and biology. The structural and non-structural proteins are described comprehensively. Additionally, available diagnostic tests and potent PPRV vaccines are discussed and finally current challenges and future possibilities for disease eradication are highlighted. Officially, PPR was first described in the Republic of Côte d’Ivoire in West Africa in 1942 (Gargadennec and Lalanne, 1942), however, there are indications that the disease existed much earlier. Since PPR and RP are clinically related diseases and the viruses are antigenically similar, it is believed that PPR remained undiagnosed due to the high prevalence of RP and the inability of the available diagnostic tests to differentiate PPR from RP (Baron et al., 2011). Furthermore, it is likely that, owing to cross-neutralization between PPRV and RPV, small ruminants infected with RPV would have developed protective antibodies suppressing the clinical outcome of PPRV infection (Taylor, 1979). Nevertheless, the disease gained attention when a severe rinderpest-like disease was observed in sheep and goats, which was unable to transmit to the cattle reared in the same herd or in the close vicinity. Initially, different names such as ‘kata’, ‘pseudo rinderpest’, ‘syndrome of stomatitis-pneumoenteritis’ and ‘ovine rinderpest’ were used to describe the disease. Later, a French name, ‘peste des petits ruminants’, was suggested because of its clinical, pathological and immunological similarities with RPV. At the time of first PPRV recognition, it was considered a variant of RPV. However, Gibbs et al. (1979) revealed that PPRV is biologically and physico-chemically distinct and is therefore a new member in the genus Morbillivirus, along with RPV, canine and phocine distemper viruses (CDV and PDV), measles virus (MV) and morbilliviruses of porpoises, dolphins and cetaceans (PMV, DMV and CMV). After first identification, PPRV spread to sub-Saharan Africa, the Middle East, Turkey and the Indian subcontinent. During the last decade, the disease has been reported for the first time in China, Kenya, Uganda, Tanzania, Morocco and Tunisia (Banyard et al., 2010; Munir et al., 2013). This demonstrates that the virus is highly infectious, and is of emerging transboundary nature. Initially, PPRV was characterized and phylogenetically analysed based on the fusion gene (F), which classified all the strains of PPRV into four distinct lineages (Shaila et al., 1996; Dhar et al., 2002). Later, it appeared that phylogenetic analysis based on the nucleoprotein gene (N) presented a better molecular epidemiological pattern (Kwiatek et al., 2007) and is currently preferred over F gene-based phylogenetic analysis. However, all the PPRV strains remained in the same group regardless of what gene was used as basis for classification, except that the F gene-based lineage I (i.e. Nig/75) became lineage II on the N gene-based tree. Recently, Balamurugan et al. (2010) suggested that the use of the haemagglutinin-neuraminidase (HN) gene, in addition to the F and N genes, could give better resolution and permit tracing of virus transmission within outbreaks. Nevertheless, it is still unclear whether differences between lineages merely reflect geographical speciation or if they are also correlated with variability in pathogenicity between isolates (Banyard et al., 2010). PPRV belonging to lineages I and II have exclusively been isolated from the countries in West Africa, where PPRV once originated. Lineage III is restricted to the Middle East and East Africa. Though lineage IV was strictly considered an Asian lineage, it is now overwhelming the other lineages in African countries, while still being predominant in Asia (Kwiatek et al., 2011; Munir et al., 2013) (Fig. 5.1). Most recent reports of PPRV in previously PPRV-free countries belong to lineage IV, which suggests that lineage IV is a novel group of PPRV and may replace the other lineages in the near future. It is also likely that only lineage IV is currently causing outbreaks. Moreover, it is crucial to note that countries once exclusively carrying a single lineage are now simultaneously reporting the presence of several lineages, i.e. Sudan and Uganda (Kwiatek et al., 2011; Luka et al., 2012). In the majority of cases, the newly introduced lineage is lineage IV (Kwiatek et al., 2011; Luka et al., 2012; Cosseddu et al., 2013). PPR is generally considered a major constraint for small ruminant production; however, the economic impact of the disease has not been fully evaluated (Ezeokoli et al., 1986; Rossiter and Taylor, 1994; Nanda et al., 1996). The economic importance of PPR is primarily due to its highly contagious nature, with a case fatality rate as high as 100%. This is of particular concern for the economics of small rural farms, where sheep and goats are reared as the sole source of income. Moreover, PPR is most prevalent in countries that rely heavily on subsistence farming of small ruminants for trade and food supply. The disease consequences can be prevented by the use of highly efficacious vaccine. It has been calculated that an investment of US$2 million can bring a return of US$24 million. This estimation has been made on 1 million animals (Stem, 1993). These facts lead to the perception that PPR is one of the top ten diseases in sheep and goats that are having a high impact on the poor rural small ruminant farmers (Perry et al., 2002). Collectively, it was estimated that PPR causes a loss of US$1.5 million annually in Nigeria (Hamdy et al., 1976), US$39 million in India (Bandyopadhyay, 2002) and at least US$1.5 million in Iran (Bazarghani et al., 2006) and US$15 million in Kenya (Thombare and Sinha, 2009). Besides these figures, the worldwide economic impact of PPR largely remains elusive, and a well-planned cost-benefit analysis of PPR versus policy responses that includes both the direct and indirect impacts associated with PPR is required. Fig. 5.1. Phylogenetic analysis of PPR isolates based on the N gene. The distribution of different lineages is detailed by shading the maps for each lineage. Like other paramyxoviruses, PPR virions are enveloped, pleomorphic particles (Fig. 5.2A) and are comprised of single-stranded RNA genome with negative polarity. The length of the entire genome of PPRV is 15,948 nucleotides, which is the second longest among all morbilliviruses after a recently characterized feline morbillivirus (Bailey et al., 2005; Woo et al., 2012). The diameter of PPR virions ranges from 400 to 500 nm. The phosphoprotein (P) acts as a co-factor of large protein (L), which is the viral RNA dependent RNA polymerase (RdRp). There are three proteins associated with the host cell membrane-derived viral envelope. The matrix (M) protein acts as a link, which associates with the nucleocapsid and the two external viral proteins, the fusion (F) protein and the HN protein. The thickness of the PPRV envelope varies from 8 to 15 nm and the length of the surface glycoproteins ranges from 8.5 to 14.5 nm (Durojaiye et al., 1985). The N protein surrounds the genomic RNA along with two other viral proteins, the L protein and the P protein to form the ribonucleo-protein (RNP). This RNP core encloses the entire genome of PPRV and protects from endonuclease digestion. The RNP strands appear as a herring bone with a thickness of ~14–23 nm (Fig. 5.2B) (Durojaiye et al., 1985). Each molecule of N protein is associated with the six nucleotides of the genome, which explains the requirement of ‘the rule of six’ for paramyxoviruses including PPRV (Lamb and Kolakofsky, 2001). Contrary to this strongly accepted belief, it was revealed that PPRV obey the rule of six but carry a degree of flexibility. By a still unknown mechanism, transcription and replication in PPRV mini-genome can accommodate some deviation in genome length, such as +1, +2 and –1 nucleotides (Bailey et al., 2007). Given the fact that the PPRV genome contains 15,948 nucleotides (multiple of six bases), 2650 copies of N proteins are required to completely wrap up the genome. Electron microscopic analysis of nucleocapsid-confirmation in other morbilliviruses (Bhella et al., 2004) indicates that approximately 13 copies of the N protein constitute a single helix, and therefore a genome would involve around 200 turns of the nucleocapsid helix. As is indicated in Fig. 5.2B, the individual cell may contain several copies of the encapsidated RNA PPR virus. Fig. 5.2. (A) Schematic structure of PPRV, along with a key to different viral proteins. (B) Uranyl acetate and lead citrate staining of the microvilli collected from experimentally infected goats. Herringbone-like cytoplasmic nucleocapsids are labelled with white arrows and the extracellular virions extruding from cytoplasm are labelled with black arrows. (Second part of the figure is reproduced from Bundza et al. (1988) with permission.) The PPRV genome carries six transcriptional units; each encodes for a contiguous and non-overlapping protein except the P gene, which also expresses C and V nonstructural proteins by an alternative open reading frame and RNA editing, respectively (Mahapatra et al., 2003). All the genes in PPRV are arranged in an order of 3′-N-P/C/V-M-F-HN-L-5′ (Bailey et al., 2005) (Fig. 5.3). An inter-genic region of variable lengths separates one gene from the other (Barrett et al., 2006). Notably, owing to variable lengths of the intergenic region between the M and F genes (without having an effect on the protein lengths), the genome varies among morbilliviruses. So far, no obvious role for this variable and high GC content intergenic region has been observed in the replication of the morbilliviruses. The sequence between two consecutive genes is AAAACTTAGGA and is highly conserved throughout the morbilliviruses, including PPRV, indicating that this stretch of sequence is important for viral replications. Data from other paramyxoviruses demonstrate that nucleotides before CTT (underlined in the intergenic sequence) indicate the end of one gene (GE) and are essential polyadenylation sites, whereas the sequence after CTT is the start of next gene (GS). It has been demonstrated that the 3′ and 5′ untranslated regions (UTRs) at both ends of the paramyxovirus genome, known as genome promoter (GP) and anti-genome promoter (AGP) respectively, are crucial for viral transcription and replication (Lamb and Kolakofsky, 2001) (Fig. 5.3). In PPRV, the 3′-genome terminus, a seat for the attachment of the RdRp polymerase complex, is a stretch of 107 nucleotides, which includes the 52-nucleotide leader region, and 3′ UTR of the N gene, both separated by a trinucleotide (GAA). This stretch of GP before the N gene’s open reading frame (ORF) start codon acts as a promoter for the synthesis of viral RNA (Bailey et al., 2007). The gene start and polyadenylation signal are located 52 nucleotides downstream of the N ORF stop in PPRV, which is highly conserved among the morbilliviruses. Recently, an in vitro transcription complex was synthesized for PPRV and it was shown that a RNP complex, collected from infected insect cells, is active in synthesizing RNA (Yunus and Shaila, 2012). As with other paramyxoviruses, the N gene present at the 3′ end of the genome was the highest transcribed gene, whereas the L gene present at the end of the genome was the least, owing to attenuation at each gene junction. Quantitative analysis indicated that the level of P mRNA synthesis is 50%, if mRNA for the N gene is taken as 100%, which means that the P gene is transcribed only once out of two attempts of polymerase. Similar analyses show that the synthesis of the L gene mRNA occurs only once out of 75 attempts. Collectively, it was shown that 50% of the total transcripts consist of N mRNA only, and the remaining 50% of all other genes (Yunus and Shaila, 2012). Using this system, it is likely that the post-transcriptional modification activities associated with the L protein of PPRV will also be explored in the near future. Fig. 5.3. Genome organization of typical PPRV. All the genes are organized in the order of 3′-N-P/C/V-M-F-HN-L-5′, flanked by 3′-genome and 5′-antigenome promoters responsible for the synthesis of + sense RNA and – sense RNA, respectively. Each of the promoters carries sequence for the leader or trailer along with sequence (non-coding region, NCR) from respective genes. Between two genes, there is a stretch of sequence divided into the gene start (GS) of first gene and gene end (GE) of the next gene. It is only P gene that encodes for C and V proteins via alternative open reading frame and RNA editing (insertion of G at the place of arrow after three Gs), respectively. The AGP, which is responsible for the synthesis of genome-sense RNA, is the complement of the 5′ UTR after the L protein stop codon, including the trailer region that becomes the 3′ end of the antigenome. The conserved 3′ and 5′ termini in the entire family reflect the similarity in their promoter activities lying in these regions. A nucleotide stretch of 23–31 at the 3′-terminus of both the GP and the AGP in PPRV is highly conserved and is considered to be an essential domain required for promoter activity. This region is believed to interact with a conserved area comprising a succession of three hexamer motifs (CNNNNN). Although the exact mechanism of domains interaction is unclear, a model has been proposed that predicts that the three hexamer motifs in the second promoter element lie on the same face of the helix, exactly above the first three hexamers at the 3′ terminus (Lamb and Kolakofsky, 2001). It is therefore more likely that these two regions in the GP and AGP interact directly with each other to form a functional promoter unit. A similar assembly is also presented in the promoters of the other paramyxoviruses (Murphy and Parks, 1999). At the junction of the GP and N gene start, a conserved intergenic triplet sequence (CTT) is also considered necessary for transcription (Mioulet et al., 2001). In an effort to construct a minigenome for PPRV, Bailey et al. (2007) demonstrated the role of GP and AGP by using chimeric mini-genomes of PPRV and RPV. They showed that the use of PPRV-AGP decreased the ability of RPV to rescue the chimeric mini-genome, which predicts the difference in closely related viruses. Moreover, it was shown that AGP is a very strong promoter and is responsible for the production of the full-length negative sense genome, whereas the GP is responsible for both transcription of virus mRNAs and transcription of the full-length positive sense virus genome. Owing to its location at the 3′ end of the genome, the gene that encodes for the N protein is the most transcribed among all genes for both structural and nonstructural proteins of PPRV. The length of the N protein of both PPRV and RPV is 525 amino acids. However, the mobility pattern on SDS-PAGE varies between different strains of PPRV and RPV. It has been observed that the N protein from African isolates (e.g. Nig/75/1) of PPRV moves faster (~55 kDa) than N proteins from the Arabian Peninsula (DORCAS_87) (60 kDa), but migrated slower than the RPV (66 kDa) (Taylor et al., 1990). Therefore, the mobility pattern was considered a biochemical marker for the differentiation of PPRV from RPV and among different strains of PPRV (Lefevre and Diallo, 1990). This difference can be attributed to the post-transcriptional modifications such as glycosylation. Now, due to the availability of the sequences, it is possible to in Silico demonstrate that the Arabian isolate DORCAS_87 only has one glycosylation site at the 65NGSK position, whereas the African strain Nig/75/1 has two glycosylation sites at 65NGSK and 444NGSE positions. This difference could contribute to the difference in the mobility of respective strains on SDS-PAGE. Moreover, it is conceivable that N protein is highly susceptible to proteolysis and degradation products may vary between the two viruses and hence differ in molecular weight. However, such predictions require experimental confirmation. The N protein plays an essential role in the replication of PPRV (Servan de Almeida et al., 2007). It has been demonstrated that silencing of the N mRNA can block the production of N transcripts and the expression of N protein. Additionally, such shutting down of the N protein indirectly inhibits the production of M protein. Collectively, the shut down of these proteins results in the inhibition of PPRV progeny by 10,000-fold (Keita et al., 2008). A region for efficient siRNA inhibition has now been identified, which is 5′-RRWYYDRNUGGUUYGRG-3′ (where R is A or G, W is A or U, Y is C or U, D is G, A or U and N is any of the four bases). Although this region is found to be common in most of the morbilliviruses, targeting a single region may not prevent the risk of escape mutants. Therefore, in case of therapeutic application of this technique, multiple targets need to be used. Besides the essential role of the N protein in viral replication and transcription, it regulates host cell protein 72 (hsp72), interferon regulatory factor 3 (IRF3) and cell surface receptors in several morbilliviruses to indirectly promote viral RNA transcription (Zhang et al., 2002; Laine et al., 2003). However, such functions are not described for the N protein of PPRV. The N protein is the most accumulated protein in infected cells and is antigenically most conserved among morbilliviruses (Libeau et al., 1995). Being most abundant, N is a highly immunogenic protein. However, the immune responses generated against the N protein are non-protective due to intra-viral location of the protein. Given its abundance and antigenic stability, the N protein has extensively been targeted for diagnostic assays (Munir et al., 2013). Apart from its diagnostic application, the genetic diversity of the N gene has been the basis for the classification of PPRV into four lineages. This classification better represents the geographical origin than the classification based on the variation of the external glycoprotein, the F protein (Diallo et al., 2007; Kwiatek et al., 2007). The phosphoprotein of PPRV, as other morbilliviruses, is acidic in nature and undergoes intensive post-translational phosphorylation (hence the acronym phosphoprotein), owing to richness in serine and threonine (Diallo et al., 1987). Due to this, the P protein migrates more slowly (79 kDa) than its predicted molecular weight (60 kDa). The phosphoprotein of PPRV (strain Turkey/00) has a high serine (Ser), threo-nine (Thr) and tyrosine (Tyr) content (Ser: 38, Thr: 8, Tyr: 5). Approximately 50% of the potential phosphorylation residues in P proteins have high prediction scores (Pred values >0.6); however, potential phosphorylation sites vary between different strains of PPRV. The length of P proteins varies from 506 to 509 amino acids between different morbillivirus members and the P protein of the PPRV is the longest among all. Despite the essential role of the P protein in viral replication and transcription, it is one of the least conserved proteins, which is demonstrated by the fact that the P proteins from PPRV and RPV share only 51.4% amino acid identity (Mahapatra et al., 2003). Moreover, the region from 21 amino acid to 306 amino acid contains the majority of uncon-served residues. Given the fact that the C-terminus of the P protein is involved in the N–P interaction, this terminus is more conserved compared with the N-terminus of the P protein. In morbilliviruses, the P protein plays crucial roles at multiple levels in both viral replication and immune regulation. For instance, the N–P interaction is required for key biological processes such as cell cycle control, transcription and translation regulation (Johansson et al., 2003). The motifs required for the interaction of RPV P protein with N protein (N–P interaction) are conserved in the P protein of PPRV. Moreover, the P protein is the vital element of the viral L–polymerase complex, and it is assumed to be a key determinant of cross-species morbillivirus pathogenicity (Yoneda et al., 2004). Despite these crucial roles of the P protein in the replication of morbilliviruses, its function in PPRV replication and pathogenicity remains elusive, which warrants future investigations. The ORF for the M protein of PPRV is located at nucleotide position 3438–4442, which is translated to a protein of 335 amino acids with a predicted molecular weight of 37.8 kDa. It is therefore considered one of the smallest proteins among all the structural proteins of morbilliviruses. The protein is highly conserved and a 92.5% and 85.0% similarity and identity have been calculated between PPRV and RPV, respectively. This high degree of conservation may reflect the essential role of the M protein in the formation of progeny viruses and interaction with the surface glycoprotein in the cell membrane. Three ATG repeats (956tctATGATGATGtca970) are identified in the gene for the M protein of PPRV, RPV and MV, while the M gene of CDV, PDV, DMV lacks this domain (Muthuchelvan et al., 2005). The M protein constitutes the inner coat of the viral envelope and acts as a bridge to connect the surface glycoprotein (F and HN) with that of ribonucleoprotein core (genome, N, P and L) (Fig. 5.2). In an effort to construct a marker vaccine candidate, it was noticed that if the M protein of RPV was replaced with the corresponding protein of PPRV, it did not affect the growth of RPV in cell culture (Mahapatra et al., 2006). Although the mechanism behind compatibility remains to be determined, it at least indicates the high level of M protein conservation among morbilliviruses. The M protein mediates the viral budding process preferentially at specialized regions of the host membrane. For instance, the budding of MV occurs at the epical microvilli in epithelial cells due to highly concentrated actin filaments, which are required for the cellular transport (Riedl et al., 2002). Electron microscopic images of intestinal epithelial cells from a goat experimentally infected with the Malig-Yemen strain of PPRV indicated that viral particles were released from the microvilli and shed in faeces (Bundza et al., 1988). Moreover, the motif (FMYL) at amino acid position 50–53 required for the localization of the M protein in the cell membrane to facilitate the budding process in Nipah virus, a member of the same family, was found to be identical with that in the M protein of PPRV (Ciancanelli and Basler, 2006). However, it is not known whether these viruses share functional homologies in their M proteins. The F protein (59.137 kDA) is one of the highly conserved proteins not only between PPRV and RPV but also among all the morbilliviruses. This conservation probably reflects the cross-protection between PPRV and RPV (Taylor and Abegunde, 1979). In all paramyxoviruses, the F protein is embedded in the viral lipid bilayer envelope and protrudes as spikes on the viral surface (Fig. 5.2). The cleavage of the F protein is a key mechanism of paramyxovirus virulence. The naïve form of the F protein (F0) undergoes post-translational proteolytic cleavage and results in two active subunits, F1 and F2. This mechanism is not well understood for PPRV. However, it has been shown that PPRV carries RRTRR at position 104–108 (Chard et al., 2008), which is recognizable by the trans-Golgi associated furin endopeptidase consistent to the cleavage site RRX1X2R (X1 indicates any amino acid, but X2 must be either arginine or lysine) proposed for the morbilliviruses. It has been shown by Rahaman et al. (2003) that the membrane-anchoring subunit of F1 of PPRV contains four well-described conserved motifs: an N-terminus fusion peptide (FP), heptade repeat 1 (HR1), HR2 and a transmembrane (TM) domain. The 3D structure of the HR1–HR2 complex has revealed that the heterodimer between HR2 and HR1 covers the inner core of the HR1 trimer, resulting in a six-helix bundle. The molecular mechanism of PPRV budding is not known, but it is likely based on identical structure of heptade repeats, which have a common fusion mechanism. It has further been shown that on anchoring the FP domain in the membrane, dimerization of the HR domains leads to fusion between the host cell membrane and the viral envelope by bringing them close to each other (Rahaman et al., 2003). A leucine zipper motif, present in the F protein of all the morbilliviruses, is responsible for facilitating the oligomerization and fusion function of the F protein through an unknown mechanism (Plemper et al., 2001). In PPRV, this motif is located at position 459–480 and is conserved among all PPRV strains characterized so far. In all morbilliviruses, the membrane-associated proteins are glycosylated and hence are known as glycoproteins. This post-transcriptional modification is critical for the transport of the protein to the cell surface, and to maintain its fusogenic ability and integrity. All members of the morbillivirus genus contain a conserved NXS/T (X indicates any amino acid) glycosylation site in the F2 subunit of the mature protein (Meyer and Diallo, 1995). In PPRV, the three N-linked glycosylation sites include 25NLS27, 57NIT59 and 63NCT65; however, their specific functions still need to be revealed. The ORF for the HN protein gene starts from 7326 and ends at 9152 nucleotide (Nigeria 75/1) and results in a 67 kDa HN protein. The HN protein is the least conserved. While both PPRV and RPV have 609 amino acid residues in their respective HN proteins, the proteins share only 50% amino acid identity. This variation probably reflects the viral specificity for cell tropism and therefore determines the host range. Most of the viral neutralizing antibodies are mainly directed against the HN protein. Hence it is under continuous increased immunological pressure (Renukaradhya et al., 2002). The fundamental roles of the HN proteins in progression of viral infection and specific binding to host cell membrane are not defined in PPRV. However, the findings that the H protein is a major determinant of cell tropism in MV and is the main cause of cross-species pathogenesis in lapinized RPV (Yoneda et al., 2002) indicate that H is the vital antigenic determinant of the morbilliviruses. However, it has been determined that the HN protein of PPRV required a homologous F protein for proper functioning in virus replication (Das et al., 2000). In some paramyxoviruses, surface proteins can cause haemagglutination and can carry neuraminidase activities. Interestingly, among morbilliviruses it is only MV and PPRV that have haemagglutination capabilities (Varsanyi et al., 1984; Seth and Shaila, 2001). In addition to haemagglutination (viral attachment to cell surfaces and agglutination of erythrocytes), PPRV is unique for its neuraminidase activity (cleaves sialic acid residues from the carbohydrate moieties of glycoproteins). Therefore, it is the only member of the morbilliviruses that has HN protein (Seth and Shaila, 2001), which was previously thought to be absent. RPV, which as already mentioned is very closely related to PPRV, has limited neuraminidase activity but cannot act as a haemagglutinating agent for the erythrocytes (Langedijk et al., 1997). Based on these results, it is suggested to use the more descriptive term HN protein instead of the currently used H protein, as has been used for PPRV in this chapter. The L protein of PPRV is 2183 amino acids long and is regarded as the largest protein in PPR virions. However, due to natural attenuation at each gene-junction in all mononegaviruses, the mRNA encoding for the L protein is the least abundant (Flanagan et al., 2000; Yunus and Shaila, 2012). Notably, the L protein is conserved among morbilliviruses: PPRV has an identity with RPV and CDV of 70.7% and 57.0%, respectively (Bailey et al., 2005). The protein is rich in leucine and isoleucine, which can be as high as 18.4% (Muthuchelvan et al., 2005). The L protein of PPRV carries a length (2183 amino acids) and molecular weight (247.3 kDa) identical to that of RPV, MV and DMV; however, the protein charge +14.5 is different from those of RPV (+22.0) and PDV (+28.0). In all morbilliviruses, the L protein acts as RNA-dependent RNA polymerase and performs transcription and replication of the viral genomic RNA. Additionally, the L protein is also responsible for capping, methylation and polyadenylation of viral mRNA. All these steps are crucial for efficient replication of the viruses. Although the direct actions of L proteins are not investigated for PPRV, it is possible to make speculations owing to high sequence identity among morbilliviruses. Three motifs in the L protein have been identified, which are directly linked to the functions of this protein. The corresponding sequences at all these sites are found to be identical in the L protein of PPRV (Munir et al., 2013). The L gene start motif (AGGAGCCAAG) in PPRV, in accordance with the motif found in other morbilliviruses [AGG(A/G)NCCA(A/G)G], is responsible for the generation of viral L gene mRNA and signal for the capping. In PPRV, the corresponding motif required for the binding of L protein with the RNA in morbilliviruses is KETGRLFAKMTYKM at amino acid position 540–553. The sequence ILYPEVHLDSPIV at positions 9–21 can act as a binding site for P and L proteins (Horikami et al., 1994). This sequence for P–L interaction is conserved in paramyxoviruses: in PPRV it is totally conserved except the first amino acid, which is valine instead (Chard et al., 2008). Although most of the important functions of L protein are not defined yet, it is expected that with the current establishment of the reconstituted system for PPRV, it will be possible to demonstrate the multifunctional activities of the L protein of PPRV (Yunus and Shaila, 2012). It is only the P gene among all the genes of PPRV that encodes for more than one protein, known as C and V proteins, through alternative open reading frame and RNA editing, respectively, only in virus-infected cells (Mahapatra et al., 2003; Barrett et al., 2006). Apart from the role of C protein in viral replication, recently it has been shown that C protein in RPV inhibits interferon beta (IFN-β) production (Boxer et al., 2009). The molecular mechanism of inhibition still needs to be investigated, but it is likely that the C protein blocks the activation of transcription factors which are required to make up the IFN-β enhanceosome. Whereas the C protein is known to be a virulence factor in MV infection (Patterson et al., 2000) and RPV growth (Baron and Barrett, 2000), the biological function of the C protein in PPRV biology is not known and needs to be examined. The length of the V protein of PPRV is highly variable among morbilliviruses (Table 5.1). The predicted molecular mass and iso-electric point of the V protein of PPRV is 32.28 kDa and 4.68, respectively. By virtue of having the same initial gene frame, the V protein shares the N-terminus to the P protein, but due to RNA editing, the cysteine-rich C-terminus is different (Mahapatra et al., 2003). The V protein, in contrast to the C protein, undergoes phosphorylation and ~60% of the serine residues are revealed to have a high score for phosphorylation as predicted by Netphos 2.0 (Blom et al., 1999). In the majority of the paramyxoviruses, the V protein antagonizes interferon actions. Studies are required to investigate the functions of the V protein of PPRV, and its relation to other morbilliviruses. My preliminary results indicate that both C and V proteins are associated with IFN regulations at ISRE level in an in vitro reporter system. However, the molecular mechanisms of inhibition might differ in both proteins (M. Munir, unpublished data). Table 5.1. Nucleotide (nt) and amino acid (aa) comparison of open reading frames between genes of PPRV (Turkey/00, AJ849636) and other morbilliviruses. RPV was first successfully grown on bovine kidney cells, but Gilbert and Monnier (1962) were also able to isolate PPRV on primary lamb kidney cells. However, later, because of the problematic quality and considerable variations in primary cultures, an African green monkey kidney (Vero) cell line was used for PPRV isolation (Lefevre and Diallo, 1990). To further improve the isolation method and to reduce the problems associated with the Vero cell line (i.e. low virus isolation, unsuccessful attempts and blind passages) (Abu Elzein et al., 1990; Lefevre and Diallo, 1990), monkey CVI cells expressing sheep–goat signaling lymphocyte activation molecule (SLAM) has been investigated. It was shown that the monkey cell line, designated CHS-20, is highly sensitive for isolation of wild-type PPRV from clinical specimens (Adombi et al., 2011). Studies have shown that SLAM can be a co-receptor for PPRV, which was first confirmed using the small interfering RNA (siRNA) technique (Pawar et al., 2008). Under silenced SLAM receptor in B95a cells (a marmoset lymphoblastoid cell line), PPRV replication was observed to be reduced by 12- to 143-fold, while the virus titre ranged from log10 1.09 to 2.28 (12–190 times). Taken together, expression and distribution of SLAM was directly proportional to that of PPRV cell tropism, indicating that SLAM may act as a receptor for PPRV infectivity. The mRNA level of SLAM was determined to be higher in lymph nodes and was detectable in the digestive system; however, despite the fact that PPRV also replicates in the lungs, colon and rectum, the SLAM receptors were not activated, which partially demonstrates that SLAM is not the major receptor for PPRV infectivity, and that PPRV additionally relies on other receptors for viral pathogenesis (Meng et al., 2011). Recently, in an experimental study it was shown that alpine goats are highly susceptible to Morocco strains of PPRV (Hammouchi et al., 2012). The results of this and a corresponding study from the same group concluded that alpine goats can be used for both vaccine and pathogenesis studies in order to consistently reproduce PPR clinical signs in experimentally infected animals (El Harrak et al., 2012). There are several factors that contribute significantly in disease pathology and virus dissemination, some contributed by the host while others are physical factors. Although only domestic and wild small ruminants are considered as the main natural host, PPRV can infect other species such as cattle, pigs, buffalo, camels, and as recently reported, also the Asiatic lion (Balamurugan et al., 2012a). There is little information available about susceptibility, occurrence and severity of the disease in wild ungulate species; however, current literature indicates that wild small ruminants may have a crucial role in the epidemiology of PPR (Munir, 2013). In small ruminants, the severity of the disease may vary depending on age, sex, breed and seasons (Amjad et al., 1996; Brindha et al., 2001; Dhar et al., 2002; Munir et al., 2009; Meng et al., 2011). Generally, it is believed that goats show more severe clinical signs than sheep in the same environmental conditions. This is supported by the fact that the level of PPRV antibodies is higher in sheep than goats, which may render sheep resistant to the disease (Munir et al., 2009). Wosu (1994) has also shown that the rate of recovery is lower in goats than in sheep. The information regarding viral preference for sheep over goats has not been investigated, but it is likely that sheep show higher natural resistance to the disease. Notably, PPRV infection can spread between goats without affecting nearby sheep (Animal Health Australia, 2009), but mixed raising of both sheep and goats is considered to be a main risk factor for seropositivity in sheep flocks (Al-Majali et al., 2008). It is also plausible that owing to the high fertility rate in goats there may be larger flock replacement by goat offspring, which are more susceptible to the disease than adults due to decrease in maternal antibodies after 4 months (Srinivas and Gopal, 1996; Ahmed et al., 2005). Furthermore, it has been demonstrated that age is the main factor for seropositivity in small ruminants (Waret-Szkuta et al., 2008). The case fatality rate is higher in young goats than in adults (Shankar et al., 1998; Atta-ur-Rahman et al., 2004). Since the males are sold earlier and females are kept for longer, the sex-based distribution of antibodies is usually biased. Goat species from West Africa are more susceptible than European goats (Couacy-Hymann et al., 2007). The dwarf varieties of goats are the most susceptible among African breeds. The disease rate (morbidity) increases with environmental stress such as confinement of animals during winter and rainy seasons (Amjad et al., 1996; Brindha et al., 2001; Dhar et al., 2002). However, the effects of environment on the occurrence of PPR are solely based on the nature of animal husbandry conditions and socio-economic status of the farm owner. Although there have been significant contributions in understanding the risk factors, the genetic marker of disease predisposition are not determined. PPRV is highly contagious and in most cases the virus is spread from infected to healthy animals via close contact (Abubakar et al., 2012). However, PPRV is commonly shed in all secretion and excretions, such as from the mouth, eye and nose, and in faeces, semen and urine. Shedding starts after approximately 10 days of pyrexia. Since the virus is also secreted in sneezing and coughing, it is likely that transmission may occur through inhalation or contact with inanimate objects. The survival of PPRV in the dam’s milk has not been investigated; however, based on its similarity with RPV, it is likely that PPRV is also secreted in milk 1–2 days before signs appear and as late as 45 days after onset of disease. It has been observed that PPRV-infected animals start virus transmission before the onset of clinical signs (Couacy-Hymann et al., 2007). However, Ezeibe et al. (2008) studied the shedding of virus during the post-recovery state of the animal, and realized that goats infected with PPRV can shed virus antigens in faeces for 11 weeks after complete recovery. Little is known about the fragility of PPRV in the external environment. Comparison with RPV is likely to be reliable because there are many features in common. Although transmission is not impossible through fomites, it is not common either, because of the short life of the virus in dry environments (above 70°C) and in acidic (>5.6) or basic (<9.6) pH. Moreover, PPRV cannot exist for a long time outside the host because of its short half-life, which is estimated to be 2.2 minutes at 56°C and 3.3 hours at 37°C (Rossiter and Taylor, 1994). No convalescent carrier or chronic form of PPR has been reported. Primarily PPRV gains entry into the host via the epithelial lining of the oral cavity, respiratory and digestive tract. Based on this, PPR is also called stomatitis pneumoenteritis complex. Due to its high lymphotropic nature, PPRV replicates in the regional lymph node after internalization. The resultant viraemia facilitates virus dissemination to the surrounding susceptible epithelial tissues of the host. Further replication of the virus in these organs leads to establishment of lesions and clinical signs. The severity of these clinical signs depends on the age, breed, body condition and innate immunity of the host and the virulence of the virus. Moreover, concurrent bacterial and parasitic infections can further aggravate the disease. Based on these factors, the clinical outcome of the disease is divided into pera-cute, acute, subacute or subclinical (Braide, 1981; Obi et al., 1983; Kulkarni et al., 1996). However, the acute form of the disease is the most common in both sheep and goats. In the acute form of the disease, a short incubation period (3–4 days) is followed by pyrexia and severe diarrhoea, which ends in emaciation and prostration. Catarrhal discharges around nostrils can lead to severe dyspnoea, sneezing and coughing (Fig. 5.4A,B). Crusting and congestion of conjunctiva at the medial canthus and conjunctival sac may eventually cause complete closure of the eyelids. Rough necrosis is common on the dental pad, hard palate, inner side of cheek and dorsal part of the tongue, and around the commissures of the mouth. Because of these lesions, animals are reluctant to open their mouth and thus become anorexic. Occasionally, lesions may also develop in the mucous membrane of vulva and the vagina in female animals, which may cause abortion in pregnant animals (Fig. 5.4C,D). The lungs are affected in PPRV-infected animals, causing dyspnoea and productive cough. Severe signs of pneumonia such as noisy respiration with extended head and neck, nostril dilation, protruded tongue and painful cough are indications of poor prognosis. The affected animals then gradually become dehydrated, with sunken eyeballs, and often die 10–12 days post-pyrexia. The case fatality rate ranges from 70 to 80%, while survivors recover after weeks of convalescence.
5.1 Introduction
5.2 PPRV Identification and Historical Perspective
5.3 Geographical Distribution
5.4 Economic Impact of PPR Disease
5.5 Virion Morphology, Structural and Accessory Proteins
5.5.1 PPR virions
5.5.2 Viral ribonucleoprotein
5.5.3 Genome organization, replication and transcription
5.5.4 Structural proteins
Nucleocapsid (N) protein
Phosphoprotein (P)
Matrix (M) protein
Fusion (F) protein
Haemagglutinin-neuraminidase (HN) protein
Large (L) protein
5.5.5 Accessory proteins
C protein and V protein
5.6 In vitro Cultures and Animal Model
5.7 Determinants of Virulence
5.7.1 Host factors
5.7.2 Non-host factors
5.8 Pathophysiology and Clinical Presentation
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