Peste Des Petits Ruminants: A First Retrospective Investigation Among Susceptible Animal Species in Qatar


 A retrospective study was conducted to investigate the prevalence of peste des petits ruminants (PPR) in Qatar. Three hundred sixty-eight blood, swabs, and organ tissue samples collected between 2009 and 2016 were screened for PPR viral antibodies, antigens and nucleic acids using ELISA-Ab, ELISA-Ag and rRT-PCR, respectively. Fifteen PPR positive samples were subjected to virus isolation using Vero cell lines. 52% (n=192) of the samples were shown positive for PPR reporting first time infection of 52% (n=71) animal species including sheep, goat, deer, gazelle, addax, Oryx, blackbuck, deer, springbuck and waterbuck. Eight PPR virus (PPRV) field isolates demonstrated classical PPRV cytopathic effect (CPE) and shown positive for the virus antigens proving finally virus isolation. Sheep had the highest infection rate (55%) followed by wild ruminants (54%) and goats (47%). History wise, PPR might exist in Qatar before 2009. A systematic investigation is recommended to identify the risk factors associated with exposure of the susceptible animals to PPR infection, to test the susceptibility of the different species to PPR infection, and to describe the molecular entity and the replicative potentiality of the circulating field strains.


Introduction
Peste des petits ruminants (PPR) is a highly infectious viral disease of domesticated small ruminants (Gibbs et al., 1979) characterized by various clinical manifestations of respiratory and digestive nature (Libeau et al., 2014) which are mostly accompanied by ulcerative in ammations and high morbidity and mortality rates (Hammouchi et al., 2012).
The etiology of the disease PPR virus (PPRV) belongs to the genus Morbillivirus under the family Praramyxoviridae (Adams et al., 2017). The virus is enveloped, non-segmented, negative sense and single-stranded. The genomes of different isolates are of variable lengths (Chard et al., 2008). However, isolates that have genes organized in the order 3' N-P/C/V-M-F-H-L 5' code for 6 structural and 2 nonstructural viral proteins (Bailey et al., 2005). Transmission of the virus is through the respiratory route.
Infection is initiated by the surface spike-like H and F structural glycoproteins to induce virus attachment and fusion, respectively (Diallo, 1990). With the aid of the signaling activation lymphocyte molecules (SLAM, CD150) and Nectin-4 (PVRL-4) surface receptors, PPRV exploits the lymphatic and epithelial pathways to establish in vivo or in vitro replication (Pawar et al., 2008;Birch et al., 2013). However, recently, a third endothelial endocytosis pathway was described where the virus replicates potentially to deregulate 146 genes 1-hour post viral attachment and entry (Yang et al., 2018). Distinct from the other members of the group, PPRV is classi ed with rinderpest virus, dolphin morbillivirus, canine distemper virus, phocid distemper virus, porpoise morbillivirus and human measles morbillivirus (Barrett et al., 1991). unknown PPR-infecting wild ruminants were also reported (Pruvot et al., 2020). Apparently healthy ruminants including Oryx (Frölich et al, 2005), cattle (Anderson and McKay, 1994 In Qatar, the total population of PPR susceptible animal species is estimated to be 1,777, 207 (DAR, 2021), 62% (n= 1,115,293) of which are sheep, 26% (n= 464,980) goats, 8% (n=135,976) camels, 3% cattle (n=45958) and 1% (n= 15,000) are wild ruminants. Due to the limited grazing areas in the country, most of the animal species are raised within mixed closures of open farming system, however, each occupies a separate unit. Of the total wild ruminants species, Oryx represents 82% (n=12,400) of the species. These are kept into 3 different separate natural reservation units supervised by the governmental Wild Ruminants Reservation Authority. The fem reminders and the rest of the wild ruminants species are owned by the private sectors and kept in the same manner described for the other animal species.
Recognized animal food importing country, the quarantine veterinary section at of the veterinary authority in Qatar is routinely checking the vaccination certi cation upon importation from PPR suspected or endemic countries. As a practice, the animal health section used to vaccinate susceptible animals against PPR using the live attenuated PPRV 75-1 Nigerian strain. Usually vaccinations are performed at the discretion of the animal owner, complementation with the animal movement regulations or response to a reported regional PPR outbreak. However, the rst incident of PPR in Qatar was reported by Banyard and Parida, 2015; based on viral RNA that had been retrieved from samples collected in 2010 from goats and deer. Nonetheless, to date, there is no single o cial national or international declaration indicating existence of PPR in Qatar.
With respect to the recent global movement towards preventing, controlling and eradicating PPR, there is a crucial need to raise the global awareness about the disease bene ting from the regulations set forth in the national feedback platforms (FAO, 2015a). Taking this into consideration, it is obvious that there are huge gaps in knowledge concerning the epidemiology of the PPR in Qatar. These gaps include the following questions: 1. Does PPR exist in Qatar? 2. And if so, what is the extent of the disease perpetuation among the susceptible domesticated and captivated wildlife hosts? 3. What are the lineages of the viruses that are circulating among the susceptible animals? 3. And nally, what are the possible factors that are potentially in uencing the disease to perpetuate among these animals?
Since PPR is on the World Organization for Animal Health (OIE) list of the noti able diseases, and it's currently a global thread to animal industry, it needs more attention. The aforementioned setbacks observed in Qatar coupled with the animal species existing in the country, the rearing pattern practiced in the country with the absence of previous vaccination records; have promoted the authors to conduct a rst retrospective investigation about the prevalence of the disease. Answers given to the previously raised questions would expect to offer baseline data on the disease and help establishment of an effective sustainable preventive and control programmes.

Materials And Methods
Samples: All samples, which represented the bulk of the material available at the Veterinary Laboratory (VL) facility, Qatar, and examined suitable for testing at the time of the study, were used in this investigation. Three hundred sixty-eight whole blood, blood sera, ocular and nasal swabs, and organ tissues, collected between 2009 and 2016 and stored between -40 o C and -80 o C were used for this retrospective investigation. The samples were derived from sheep, goats and wild ruminants, which had been examined by the relevant veterinary clinics (VC) upon suspecting PPR and/or foot-and-mouth disease (FMD) and blue tongue (BT). Clinical data of the samples and the putative factors that might account for the risks of exposure of these animals to PPR including duration of the sample acquisition, vaccination record, animal focal point (FP) and VC were obtained from each relevant sample submission sheet and recorded. Procedures, which had been followed to submit the samples from the VC to the VL, were traced. Brie y, each sample package had been received by the Sample Registration O ce (SRO), checked for quality and data relevance before submission to the designated diagnostic unit. Necropsy samples and the relevant data, which had been collected at the VL by the pathologist were submitted to the SRO before redirected to the diagnostic units. Whole blood and blood serum samples, which had been obtained from the life or necropsied animals, were collected each into 5 mL to 8 mL volume by venipuncture of the external jugular vein or by heart puncture, respectively. Blood sera obtained into 2 mL aliquots had been prepared in the relevant VL facility following overnight standing between 18 o C and 26 o C for coagulation before centrifuged at 800 g for 10min at 10 o C.
Enzyme-linked immunosorbent antibody: The PPRC VER1209 GB competitive enzyme-linked immunosorbent assay (cELISA) kit was used to determine the anti-PPRV Ab in each 25-µL-serum sample following the manufacture's instructions (IDvet Innovative Diagnostic Kits, Grabels, France). An in-lab preprepared and validated internal positive quality control serum was included in each run. Optical densities were obtained using BioTek ELx808 spectrophotometric microplate ELISA reader aided with a 450 nm WL lter. Results were interpreted using Gen5.5 software version and a validated ELISA-Ab spreadsheet results interpretation calculator.
Enzyme-linked immunosorbent antigen: The PPRAG 0313-506 VER0313 GB capture enzyme-linked immunosorbent assay (capELISA) was used to determine the PPR viral Ag in each swab and organ tissue collected for this purpose. About 10% of the viral swab elution and tissue homogenate concentration were used to test each sample following the manufacture's instructions (IDvet Innovative Diagnostic Kits, Grabels, France). Optical densities were obtained using BioTek ELx808 spectrophotometric microplate ELISA reader aided with a 450 nm WL lter. Results were interpreted using Gen5.5 software version and a validated ELISA-Ab spreadsheet results interpretation calculator.
Determination of PPRV cDNA Viral RNA extraction: Swab elution and blood samples (140 μL each) were used for to extract the PPRV RNA using the QIAamp Viral RNA Mini 2010 methodology. About 25-30 mg of each previously collected organ tissue was used to purify the viral RNA following the Qiagen RNeasy® Mini Kit instructions (Qiagen, Hilden, Germany). Finally, each viral RNA extract was obtained in 80-μL amounts using the kit AVE buffer (Qiagen, Hilden, Germany) as elution buffer. All aliquots were stored at -20°C till further processing.
Real-time polymerase chain reaction (rRT-PCR): The single-step qualitative real-time reverse transcription polymerase chain reaction (rRT-PCR) described by Qiagen, GMbH, Germany, 2011 was used to detect the PPRV RNA following the manufacturer's instructions. About 100 ng (10 uL) each RNA template extract was used for ampli cation to obtain the viral cDNA. Positive and negative controls were treated the same using an approximation of 0.1% PPR positive control preparation and nuclease-free water (NTC), respectively. Using ABI 7300 Real Time Cycler, a program of 48 °C for 10 min (RT step), 95 °C for 10 min (RT inactivation/denaturation), 40 cycles of 95 °C for 15 s (denaturation) and 60 °C for 45 s (annealing/extension), respectively, was used for ampli cation of the obtained cDNA fragments.
Samples selection and preparation: Based on the chronology of the date of sample acquisition, sample quality, nature of the clinical and necropsy ndings and positivity to ELISA-Ag and/or rRT-PCR, 15 tissue samples representing 8 species were selected for virus isolation into Vero cell lines. 10% homogenate concentrations were prepared using sterile phosphate-buffered saline (PBS). All samples underwent ltration using sterile 0.2 μm Millipore lters. Each nal 2 mL sample aliquot was supplemented with 100 iu/mL penicillin, 20 μg/mL streptomycin and 20 μg/mL gentamycin, incubated for 1 h at room temperature, and preserved at 4°C before inoculation.
Virus cultivation: Each 500 μL pre-prepared PPRV inoculum was inoculated onto 4-day-old 70-80% semicon uent 25-cm 3 Vero cell line following thrice washing of the cells with 5 mL PBS. All infected cultures were incubated for 1 h at 37°C into CO 2 -free cell culture grade incubator. Control asks received serumfree DMEM were treated the same. Following this, each inoculum was removed from the infected and control asks. Finally, each ask received 5 mL 2% BCS-supplemented DMEM. All asks were incubated back at 37°C and guarded daily for change in media colour and development of cytopathic effect (CPE).

Results validation
Samples from animals have had classical clinical signs and/or typical necropsy features and detected positive for PPRV Ag and PPRV RNA or succeeded virus isolation were considered PPR infected. Moreover, samples from animals have had classical clinical signs and/or typical necropsy features and detected positive for PPRV Ab were considered PPR infected. Samples from animals with no clinical features or atypical PPR clinical signs and/or postmortem ndings and detected positive or negative for PPRV Ab were considered PPR negative.

Statistical analysis
The proportion of the PPR samples considered positive for the PPR infection and indicated for the positive animals was calculated out of the total PPR-suspected samples. Previous records of the putative factors that might represent risks exacerbating exposure to the PPR-infected cases were evaluated. The signi cance of association of each factor to PPR infection was evaluated as the univariate logistic regression. All signi cances were considered at type I error of α < 0.10.
Detection of PPR viral RNA: 51% (n=170) molecularly tested samples retrieved viral RNA fragments (Table  1). Except to samples from blackbuck (PPRV-QBB14), all rRT-PCR results were on cohort with the viral isolation ndings. Study results: The 368-subject samples were collected from 135 PPR-suspected cases, 31 of which were sheep, 36 goats and 86 were wild ruminants ( Table 2). Initial screening to detect PPR Ab, Ag, nucleic acid or to isolate the virus has resulted in 57% (n= 211) positive PPR samples; 47% sample from sheep, 61% from goats and 51% from wild ruminants. Validations of the results quali ed 52% (n=192) of the screened samples positive for PPR infection and were used to identify the relevant PPR-infected animals ( Table 2). 52% (n=71) of the animals were considered infected with the eld PPRV strains, 54% (n=17) were sheep, 47% (n=17) goats and 54% (n=37) were wild ruminants. Including sheep and goats, 7 wild ruminant animal species namely; deer, gazelle, addax, Oryx, blackbuck, springbuck and waterbuck (data not shown) were considered positive for PPR infection.

Discussion
In this study, conventional and molecular assays were used to retrospectively investigate the prevalence of PPR in Qatar using samples collected from animals that have been raised between 2009 and 2016. The initial ndings con rmed existence of PPR in Qatar with a rate of 52% of the investigated animals being infected with the virus. Further, that 9 different animal species were shown infected with circulating PPRV eld isolates re ects the extent of the perpetuation of the disease among the subject animals and assumes endemic status of PPR in Qatar. This observation could be inferred from the facts that: 1. There had been continuous infections throughout the years of collection of the samples. 2. There had been repeated occurrence of PPR infection in 8% of the investigated focal points. 3. And, there had been continuous exposure to the virus veri ed by seropositivity of some of the unvaccinated animals.
The observational nding that the sample collection date of the sheep isolate PPRV-QS09 was 2009 (data not shown) de es Banyard and Parida, 2015 report that the rst Qatari PPR case was in 2010. Considering the earlier regional reports of existence of PPR evidenced by Furley et al., 1987;Hedger et al., 1980;and Taylor et al., 1990, and the infections in neighboring countries documented by Abu- Elzein et al., 1990 andElzein et al., 2004, it is likely that the disease was introduced prior to 2009 as observed in this investigation. This is probably to happen due to transmission from infected livestock imports, which have been kept in close quarters to susceptible animals. The keeping of camels in some neighboring PPRendemic countries to reduce overstocking practiced in Qatar might also increase the likelihood to expose to circulating eld strains. Failure to detect such infected cases in the meantime might be due to lack of The study observation that 7 wild ungulates were found infected with the PPRV, some of which are reported for the rst time, also indicates the wide perpetuation of the disease among these species. One consideration in this study is that gazelle and deer had higher morbidity rate (46%) compared to the other members of the species category (data not shown). This might be due to population biasness, the large number of the tested samples or due to potential moving of these animals for trade purposes subjecting these animal species to stresses. Since 54% of the PPR-infected cases were demonstrated to occur in summer (P≤ 0.001), rearing of susceptible unvaccinated animals in an open housing system, PPR endemic conditions, temperatures ranging between 40.7 o C and 45.2 o C and relative humidity between 41% and 55% might enhances sensitivity to PPR infection. The relatively large number of the involved wild ruminants species in this study correlates with the previous reports indicating expansion of PPR among new wild ruminant members (Pruvot et al., 2020), support the call to investigate the potentiality of PPR infection among wild ruminants (Al-Naeem et al., 2000), and encourages the recommendations to investigate the role of wild ruminants in the epidemiology of PPR and the status of the disease at the wildlife-livestock interface (Fine et al., 2020).
Additionally, camels, which are estimated to comprise 8% of the total PPR susceptible animals in Qatar, should be closely monitored. Since the dromedary camels play an important role in the traditional animal industry, additional investigations are required to determine whether they are reservoir hosts perpetuating PPR (Khalafalla et al., 2010) or atypical dead-end PPR-infected hosts maintaining non-infectious viral RNA replication (Schulz et al., 2019) and demonstrating serum viral Ab (Ismail et al., 1992;Roger et al., 2001;Haroun et al., 2002, Abraham et al., 2005. In regards to goats, they are considered to be most sensitive to PPR infection among domesticated species (Lefevre and Diallo, 1990), attributing this to the abundance of the SLAM cell receptors mediating the lymphoid pathway of the virus infection (Prajapati et al., 2019). Sustained by the previous observations of Mahapatra et al (2006) ndings, Pope et al (2013) concluded that upon experimentally respiratory infecting gaots, PPRV replicates in the local tonsillar and lymphoid tissues before undergoing epithelial replication, a nding concluded by early demonstration of the viral antigens at these sites and the late-mild mucosal demonstrated form of the disease observed among these animals. However, based on the clinical records, our investigation has observed some variations in this sensitivity. Higher infection rates and profound respiratory features were observed among the infected sheep and wild ruminants compared to goats (data not shown). This observation is consistent with the recent reports of an endothelial endocytosis pathway for PPRV infection (Yang et al., 2018), and agrees with the suggestion that mortalities among PPR-infected animals are due to viral pneumonitis. In our view, this logic since the respiratory system is the primary predilection site for PPR infection, rendering the lungs highly susceptible to infection due to its exceptional abundant endothelial cells lining this highly vascularized organ. Diop et al (2005) and Couacy-Hymann et al (2007b), had already reported an impact of breed, and nutritional and health status in virulence among PPRV infections. Further systematic investigation is required to identify the effect of species and breed for sensitivity to PPRV infection. Furthermore, factors including the animal of choice, sample quality (sampling duration, sample of choice, sample number, sample treatment) and reporting system should also be monitored in testing sensitivity to PPR infection among different species during natural infections and outbreaks.
The importance of PPR worldwide has obligated implementation of robust surveillance programmes to continuously guard against the disease. Rapid diagnosis using the most reliable and accurate diagnostic tools is the key factor to reach conclusive results. In this investigation we used ELISA Ag, rRT-PCR and virus isolation to differentiate PPR-infected samples from non-infected. Although others advanced techniques are also used, the results obtained are highly compatible. However, the mismatched result for the blackbuck samples isolating the virus (PPRV-QBB14) versus failure to retrieve the virus RNA is an example for seldom-unexpected results. Contamination with RNA inhibitors is common, and it might be the cause for the false negative result we obatined. Despite no speci c RNA inhibitor is known to effectively degrade PPRV RNA, exogenous and endogenous RNases of A, B and C types are usually the ascribed degrading agents compromise the quality of the PCR assays. The pancreatic ribonucleases, group of the RNase A (RNase 1), are the most endogenous tissue-sourced RNA inhibitors encountered during molecular assays, and they might be the cause for our negative rRT-PCR result. Similar to MMLV and Taq DNA polymerase, heparin, the common used blood anticoagulant, co-puri es with RNA (Jung et al., 1997) during isolation of RNA from blood, plasma, serum, buffy coat and bone marrow samples; and degrade the viral RNA (Beutler et al., 1990;Bai et al., 2000). One additional possible limitation to our result pertains to sample quality. This could be due to the long storage period of the sample prior to conduct testing (Rainen et al., 2005). Notably, the successful virus isolation was achieved from the liver sample kept as a separate set and experienced optimum storage conditions at the virology unit. Unfortunately, the investigation did not subject this sample to rRT-PCR to verify this assumption.
The continuous high rate of spread of PPR around the world is quite evident (Dou et al., 2020). This might be due to the 3-dimensional pathways of replication the virus owns enabling it to potentially infect wide range of hosts (Aziz et al., 2020) and to a ict an estimate of 30 million animal globally every year (OIE, 2018). These observations might also imply either evolution of new genetically modi ed strains that compromise the e ciency of the current applied vaccines (Mumford, 2007) or indicates gaps in the controlling programmes rendering infected animals introduced inattentively in PPR-free territories (Dormitzer et al., 2011;Tussche et al., 2012). With the recent tendency to establish national food safety and security chain in Qatar, there is a need to undertake policies to implement effective preventive and control programmes against PPR. Endorsement of regular active surveillance programmes coupled with systematic vaccination schedules is a must. The use of effective diagnostic methods and analytic tools including next generation molecular sequencing and virus isolation are advised to ensue infectivity and reveal virus phylogeny. Adoption of pro ciency testing with reference laboratories is important to build the diagnostic capacities and to ensue e ciency and certainty.