Abari, John Addra, Oladele, Sunday Blessing, Sai’du, Lawal, Orakpoghenor, Ochuko, Enam, Samson James, Elijah, John Oluwamayokun, Bilbonga, Garleya, Elijah, Mary Oluwatomisin, Ogala, Ojonigo, Sanka, Jibrin Ibrahim: Newcastle disease virus strain XIV.2 infection in geese and Guinea fowls: A comparative pathological study
ABSTRACT
Aim:
The study evaluates the pathogenic potential of strain XIV.2 Newcastle disease virus (NDV) in geese and guinea fowls experimentally.
Methods:
Forty birds, comprising 20 geese and 20 guinea fowls, aged 7 weeks were randomly divided into 4 groups (A–D), kept in separate pens as follows: 10 Control geese, 10 Infected geese, 10 Control guinea fowls, and 10 Infected guinea fowls. The birds in groups B and D were inoculated with 0.2 ml of strain XIV.2 NDV inoculum (titer of 108.2/0.1 ml ELD50) intra-nasally, while those in groups A and C were inoculated with phosphate-buffered solution intra-nasally, respectively. Gross and histopathological procedures were performed using standard methods.
Results:
Revealed transient clinical signs without mortality in infected geese whereas moderate clinical signs with mortality were observed in infected guinea fowls. Gross lesions were mild in infected geese from 6 to 28 days post-inoculation, (dpi) and in infected guinea fowls at 3, 21, and 28 dpi, with moderate lesions observed from 6, 11, 12, and 14 dpi. Histopathological lesions in infected geese were moderate from 3 to 21 dpi, but mild at 28 dpi, whereas mild lesions were observed from 3 to 6 dpi, followed by moderate lesions from 9 to 28 dpi in infected guinea fowls.
Conclusion:
Morbidity and mortality due to strain XIV.2 NDV infection were moderate in guinea fowls, but inapparent and transient in geese. Therefore, free-ranging systems commonly practiced in the region should either be discouraged or vaccination of these species of birds should be encouraged.
KEYWORDS Newcastle disease virus; pathogenesis, strain XIV.2; virulence; geese; guinea fowls
Introduction
Newcastle disease virus (NDV) is one of the most lethal poultry viral agents threatening the poultry production industries, causing Newcastle disease (ND) in domestic, cage, and wild birds irrespective of their age and sex, although some species of wild and semi-domesticated birds such as waterfowls are less susceptible but may serve as reservoir hosts [ 1]. The disease is still one of the most devastating viral diseases of poultry worldwide, causing major economic impact due to high morbidity and mortality rates as well as trade restrictions [ 2– 4]. Since the documented outbreaks of ND in the 1920s in Java, Indonesia, and Newcastle-upon-Tyne, England, it has been reported in Over 200 avian species with a diverse range of clinical outcomes [ 5]. In Nigeria, the disease was first reported in Ibadan by Hill et al. [ 6], since then it has become the most important viral disease of chickens and spread throughout the country with annual epidemics being recorded in highly susceptible poultry flocks [ 7]. The high genetic diversity of the viral strains may be responsible for the increasing rate of the disease [ 8]. The virulence of the viral strains varies significantly with the host, although breed does not seem to be the major determining factor of chicken susceptibility to the disease [ 5]. In other avian species, the disease produced by virulent ND viruses ranges clinically from inapparent to a fatal condition. The variation in severity of the disease and signs observed is mostly determined by the virus strain, the species of bird, immunity, age, and rearing conditions [ 9– 10]. Waterfowls such as ducks and geese may become infected with the virus but often show few or no clinical signs even with strains of NDV that are virulent to chickens [ 11, 12], despite evidence showing that all avian species are susceptible to NDV, including cormorants, pigeons, chickens, turkeys, parrots, migratory waterfowls, penguins, guinea fowls, and shorebirds [ 13]. Virulent strains of NDV have been isolated from double-crested cormorants, and other isolates of low virulence have been identified in different water bird species such as ducks and geese [ 14]. Guinea fowls are reported to be disease resistant, Boko et al. [ 15] stated that they are affected by colibacillosis, salmonellosis, and ND, even though chickens seem to be more susceptible to ND than guinea fowls. Moreover, a natural outbreak of ND in a flock of guinea fowls in Nigeria has been reported [ 16]. In Nigeria, like many other third-world countries, free-ranging chickens, backyard poultry, and some commercial poultry farms have no borders or measures protecting them against NDV transmission to or from semi-domesticated and wild birds [ 17]. Poultry health is of high concern in Nigeria, thus, NDV study currently focused more on domesticated species than on semi-domesticated and wild birds. Viral and microbial bidirectional spillover between semi-domesticated, wildlife, and domesticated species is now recognized but the pathogenesis of vNDV such as XIV.2 in geese and guinea fowls commonly found in close contact with backyard and commercial poultry is understudied in the region, despite the detection and isolation of genotype XIV.2 viruses in vaccinated flocks [ 18]. Therefore, this study evaluates the pathogenesis of strain XIV.2 NDV on geese and guinea fowls to understand the dynamics of the disease beyond the rural and commercial domesticated chickens.
Materials and Methods
Experimental birds
Forty birds comprising 20 geese and 20 guinea fowls at 3 weeks old were acquired from reputable breeders and raised for an additional 4 weeks to attain 7 weeks of age, screened and confirmed seronegative for NDV using hemagglutination inhibition (HI) test. The birds were kept on a deep litter system of management, fed with broiler starter (Ultima®) and cabbage (geese only) throughout the experimental period, the feed and water were supplied ad libitum.
Ethical approval
The research related to animal use complied with all the relevant national regulations and was approved by the Ahmadu Bello University Committee on Animal Use and Care (ABUCAUC) with reference number: ABUCAUC/2021/076.
The challenge virus
The NDV (XIV.2) strain biologically characterized by Welch et al. [ 8] obtained from the National Veterinary Research Institute (NVRI), Vom-Plateau State, Nigeria, was used to challenge the experimental birds, after testing its viability embryonically, with the embryonic lethal dose (ELD50) established at 10 8.2/0.1 m ELD50/ml.
Testing the viability of the challenge virus
The viability of the viral strain was tested in the Virology Laboratory Unit of NVRI, Vom as follows: 1 ml of the test sample mixed with some amount of antibiotics (penicillin and gentamycin) was inoculated into 9-day-old embryonated minimum pathogen-free chicken eggs obtained from NVRI, Vom using a sterile 2 ml-syringe and needle. The inoculation was done through the allantoic route incubated at 37°C and candled with a 25-voltage bulb daily for 4 days. Eggs that showed embryonic death within or in less than 24 hours appeared to contain viable virus. The allantoic fluid from such eggs was harvested and a confirmatory test using HI was carried out as stated by the pathogenicity test index. The titer to be inoculated into each bird (ELD50/ml) was also determined using an HI test with 1% washed chicken red blood cell (RBC) and arrived at a titer of 108.2/0.1 ml/bird as the (ELD50/ml) to be inoculated intranasally. It was then kept in vials of 1 ml at −20°C.
Experimental design
The geese and guinea fowls were randomly divided into four groups (A-D) using the random sampling method as follows: 10 control geese, 10 infected geese, 10 control guinea fowls, and 10 infected guinea fowls, respectively. Each group was kept in a separate pen, located far apart from each other. The control groups were inoculated with 0.2 ml of phosphate buffer solution (PBS) whereas the infected groups were inoculated with 0.20 ml of the viral inoculum containing strain XIV.2 NDV (Nigerian strain) with a titer of 108.2 ELD50/ml. All birds were provided with feed and water ad libitum. After inoculation, the birds were monitored for clinical signs.
Experimental groupings
| Group |
Bird composition |
Treatment |
| A |
Control geese |
Untreated |
| B |
Infected geese |
0.2 ml of 108.2 ELD50/ml titre of XIV.2 NDV inoculum intra-nasal route |
| C |
Control guinea fowls |
Untreated |
| D |
Infected guinea fowls |
0.2 ml of 108.2 ELD50/ml titre of XIV.2 NDV inoculum intra-nasal route |
NDV=Newcastle Disease Virus, ELD50=Embryonic Lethal Dose.
Serological test
Following inoculation hemagglutinin inhibition test was conducted by conventional microtiter methods as described by Allan and Gough cited by Hossain et al. [ 19] to ensure the infection had been established in the infected groups, respectively. Blood samples were collected from all the birds in each group at day 0 (pre-inoculation) and day 3, 6, 9, 12, 15, and 18 post-inoculation, respectively, into plain sample bottles without anticoagulant were allowed to clot, the sera were decanted into serum vials and frozen at −20°C. The sera were heat-treated at 56°C for 30 minutes and tested using a microtiter HI plate test with PBS as diluent. After two-fold serial dilutions of the sera were made in 96 well plates with the addition of NDV-LaSota used as antigen, it was then incubated for 30 minutes at room temperature. Washed chicken RBCs were added and the test was read after an additional 30–45 minutes. Antibody titers were expressed as the reciprocal of the dilution of sera at which there was complete inhibition.
Clinical signs
The birds in the infected groups were observed three times (visual examination) daily (in the morning, afternoon, and evening) for clinical signs of ND. The presence of clinical signs, such as anorexia, depression, huddling, ruffled feathers, whitish or greenish diarrhea, torticollis/opisthotonos, tremor, and death were carefully looked for and those found or seen were recorded with the percentage morbidity and mortality determined using the equation below.
Gross examination
Following inoculation, at 3, 6, 9, 14, 21, and 28 days post-inoculation (dpi), one live goose and guinea fowl from each of the groups were humanely euthanized and examined for gross lesions. The organs (thymus, spleen, bursa of Fabricius, intestine, proventriculus, kidney, and brain) of each bird were examined. Guinea fowls that died post-inoculation were necropsied and examined for gross lesions of NDV.
Histopathology examination
Tissue samples from the thymus, spleen, bursa of Fabricius, intestine, proventriculus, kidney, and brain were fixed in 10% neutral buffered formalin and processed for histopathology using the method of Baker et al. [ 20]. Slides were viewed using light microscope at various magnifications and the lesions were photographed using Amscope MT of the Department of Human Anatomy, College of Medicine, Ahmadu Bello University, Zaria.
Data analyses
Data were presented using Tables, Photographs, and Photomicrographs.
Table 1.Show the percentage of clinical signs in infected geese and guinea fowls from days 4–15 post-inoculation with strain XIV.2 NDV.
| Clinical signs |
Infected Geese (%) |
Infected Guinea fowls (%) |
Days-post-inoculation |
| Anorexia |
20 |
50 |
4–15 dpi |
| Depression |
40 |
60 |
4–15 dpi |
| Huddling |
0 |
80 |
4–15 dpi |
| Ruffled feathers |
0 |
70 |
6–12 dpi |
| Diarrhoea |
60 |
80 |
6–15 dpi |
| Muscular tremor |
0 |
10 |
9–11 dpi |
| Opisthotonos |
0 |
10 |
9–12 dpi |
| Death |
0 |
20 |
11–12 dpi |
Table 2.Shows the gross lesions in infected geese and guinea fowl post-inoculation with strain XIV.2 NDV.
| Days post-inoculation |
Infected geese |
Infected guinea fowls |
| 3dpi |
No visible gross lesions |
No visible gross lesions |
| 6dpi |
Mild petechiation of the thymic lobes, congested spleen and lungs, ecchymotic hemorrhages of the brain. |
Enlarged and white stippling spleen, pinpoint haemorrhages of the duodenum |
| 9dpi |
The same lesion observed as in 6dpi above. |
Atrophied thymus |
| 11 and 12dpi |
------------------------------------------ |
Hemorrhagic, congested and atrophied thymic lobes, atrophied bursa of Fabricius, congested liver and engorged gall bladder |
| 14dpi |
Heamorrhagic thymus, enlarged and congested kidneys with engorged intralobular vein |
Atrophied bursa of Fabricius and congested kidneys |
| 21dpi |
Atrophied spleen and bursa of Fabricius |
Pin-point hemorrhages of the cecal tonsils and the brain |
| 28dpi |
slightly atrophied spleen and bursa of Fabricius |
Slightly atrophied spleen and bursa of Fabricius. |
Note: Two infected guinea fowls died on days 11 and 12 post-inoculation.
Results
Clinical signs
The clinical signs observed in the infected geese were transient depression, anorexia, and white to greenish diarrhea ( Table 1). In infected guinea fowls the clinical signs manifested included depression, anorexia, ruffled feathers, whitish to greenish diarrhea, muscular tremor, opisthotonos, and death ( Table 1).
Gross lesions
There was no gross lesion in control geese and guinea fowls. Gross pathological changes observed in infected geese and guinea fowls from days 6 to 28 pi are shown below ( Table 2) and ( Figs. 1– 6).
Histopathology findings
The histopathological changes in infected geese and guinea fowls are presented below ( Table 3) and ( Figs. 7– 13).
Discussion
In this study, insignificant clinical signs without mortality were observed in infected geese, whereas moderate clinical signs with mortality were observed in the infected guinea fowls. The susceptibility and subsequent manifestation of the clinical signs in geese and guinea fowls following infection with strain XIV.2 NDV may be attributed to the presence of host cellular proteases. These ubiquitous proteases exist in a majority of host tissues, and for infection to occur, these enzymes activate and split (cleave) the viral fusion (F) glycoproteins precursor (FO protein) into two subunits F1 and F2. These proteins play a crucial role in the F of the viral genome to the host genome and the subsequent spread of the virus to different organs of the host [ 21]. Therefore, the mild clinical signs followed by insignificant morbidity and lack of mortality in the inoculated geese could result from the presence of very low cellular and tissue proteases in this species compared to guinea fowls. Moreover, genetic resistance to NDV has been observed within various breeds of chickens [ 22], turkeys, and waterfowls [ 23], and each strain of the NDV virus may be better adapted to grow in one species than another, like what was observed with PPMV-1 (pigeon) NDV strain in chicken [ 24]. Thus, the inapparent infection observed in infected geese following inoculation with strain XIV.2 may be responsible for consistent outbreaks and isolation of strain XIV.2 NDV in vaccinated chickens in the region, due to the possibility of this species harboring and perpetuating the viral strain in the area shedding through feces without manifesting clinical signs. Most of the households in the studied region reared guinea fowls as an alternative to poultry because they are relatively less expensive to raise and considerably resistant to disease compared to chicken, while geese are raised as ornamental and security birds. This increases the risks of free-ranging chicken exposure to the viral agent.
Figure 1.
Show the thymus in control geese (a) and guinea fowls (b), atrophied and petechial hemorrhages of the thymus in infected geese (c), atrophied and congested thymus in infected guinea fowls (d). 
Figure 2.
Show the spleen in control geese (a) and guinea fowls (b), enlarged and congested spleen in infected geese (c), edematous and white stippling spleen in infected guinea fowls (d). 
The gross changes observed in inoculated geese and guinea fowls were consistent with the findings of other researchers [ 25– 27]. These gross lesions observed in various organs may be attributed to the viral activities within these organs since researchers’ reports have indicated the ability of hemagglutinin neuramidase protein of NDV to play a major role in the tissue and organ tropism facilitated by the membrane F protein to enable the viral agent to gain entrance into the host cells, tissues, and organs, respectively [ 28]. Inside the host organs, the virus causes direct destruction of the endothelium and blood vessels that may stimulate hemostatic plug formation structured by thrombocytes [ 29]. These cells are present in the blood of all non-mammalian vertebrates derived from mononuclear precursor cells in the bone marrow (thromboblasts) and also have phagocytic properties [ 30]. Unfortunately, previous research reports have indicated that the acute phase or stages of NDV infection is mostly accompanied by thrombocytopenia and endothelial damage resulting in hemorrhages observed in various organs [ 30]. Therefore, the hemorrhages observed in the heart, brain, and thymus may be due to the direct viral destruction of the vascular endothelium, bone marrow aplasia, or the ability of the virus to alter the coagulation cascade that resulted in disseminated intravascular coagulopathy due to lack of clotting factors in the systemic circulation as reported by researchers in viral infections of avian species such as avian influenza, avian infectious anemia, NDV and infectious bursa disease virus [ 30]. The enlarged and multifocal white stippling spleen may be due to the depletion of splenic T-lymphocytes and subsequent increased infiltration of macrophages that induce apoptosis and increase secretion of Th1-like proinflammatory cytokines [ 31]. There was no obvious gross lesion observed in the proventriculus and gizzard of the infected geese and guinea fowls, respectively, these affirm the study of Mishra et al. [ 32] who reported a lack of specific gross changes in the proventriculus and gizzard of both the dead and euthanized guinea fowls following exposure to NDV. This absence of gross changes in these organs may be probably due to the fact that NDV strains differ considerably in the organ and system they affect, and the severity of the clinical signs manifested in infected hosts [ 33]. The histopathological changes in the thymus, spleen, intestine, proventriculus, kidney, bursa of Fabricius, and brain were consistent with the previous reports on ND in various avian species by other researchers [ 34– 38].
Figure 3.
Show intestines of control (a) and infected geese (b), control (c) and infected (d, e) guinea fowls Note: Absence of gross lesion in the duodenum of infected geese (b), petechial hemorrhages (arrows), yellowish mucoid shred (arrows head) in the duodenum and cecal tonsils in infected guinea fowls (d, e). 
Figure 4.
Shows the proventriculus of infected geese (a) and guinea fowls (b) without specific gross lesions in both infected geese and guinea fowls, respectively. 
Figure 5.
Shows kidney in control geese (a) and guinea fowls (b), enlarged kidney with congested interlobular renal veins (arrows) in infected geese (c), congested kidney in infected guinea fowls (d). 
Figure 6.
Shows the brain in control geese (a) and guinea fowls (b), congested and hemorrhagic brain in infected geese (c) and guinea fowls (d), respectively. 
Table 3.Shows the histopathological lesions in infected geese and guinea fowls post-inoculation with strain XIV.2 NDV.
| Days-post-inoculation |
Infected geese |
Infected guinea fowls |
| 3dpi |
Mild fragmentation of the intestinal villi and congested lung parabronchus. |
Mild fragmentation of the intestinal villi. |
| 6dpi |
Mild depletion of lymphoid organs |
Mild depletion of lymphoid organs |
| 9dpi and 14dpi |
Fragmentation, necrosis, and desquamated villi and proventricular glands, depletion of thymus and splenic white pulp, congested brain blood vessel and renal vein and renal vein. |
Fragmentation and desquamated intestinal villi and proventricular gland, depletion of splenic white pulp, congested brain blood vessel |
| 21dpi |
Widen interfollicular space, depletion of lymphoid organs, infiltration of mononu-clear cells around the portal area. |
Widen interfollicular space, depletion of lymphoid organs, |
| 28dpi |
Widen and thickened interfollicular space with cystic follicles |
Widen and thin interfollicular space |
Figure 7.
Photomicrographs of a section of thymus in control geese (a) and guinea fowls (b), congested (arrows) thymus in infected geese (c) and guinea fowls (d) H & E X400. 
Figure 8.
Photomicrographs of a section of spleen in control geese (a) and guinea fowls (b), depletion of splenic white pulps (arrows) in infected geese (c) and guinea fowls (d) H & E X400. 
Figure 9.
Photomicrographs of a section of the intestine in control geese (a) and guinea fowls (b), necrosis (arrow), fragmentation (blue arrow), and desquamation (arrowheads) of the intestinal villi into the lumen in infected geese (c), and guinea fowls (d) H & E X400. 
Figure 10.
Photomicrographs of a section of proventriculi in control geese (a) and guinea fowls (b), necrosis (arrows) and desquamated materials (arrowhead) into the proventricular gland in infected geese (c), edema (blue arrow), and desquamated proventricular gland (arrowhead) in infected guinea fowls (d) H & E X400. 
Figure 11.
Photomicrographs of a section of kidney in control geese (a) and guinea fowls (b), congested renal veins (arrows) and necrotized renal tubules (arrowheads) in infected geese (c), and guinea fowls (d) H & E X400. 
Figure 12.
Photomicrographs of a section of bursa of Fabricius in control geese (a) and guinea fowls (b), thickened (arrows) and widened bursa interfollicular spaces (arrows head) in infected geese (c), and guinea fowls (d) H & E X400. 
Figure 13.
Photomicrographs of a section of the brain in control geese (a) and guinea fowls (b), congested brain (arrows) in infected geese (a) and guinea fowls (b) H & E X400. 
Conclusion
Although obvious mortality caused by the viral strain in the infected birds was only observed in guinea fowls, the virus was able to induce gross and histopathological changes in all the infected birds. Therefore, the viral strain poses a significant threat to these species and the rural chickens commonly reared together, since the strain is endemic in the studied region. Therefore, poultry farmers who rear geese and guinea fowls concurrently with chickens should be encouraged to vaccinate them against NDV.
Acknowledgment
The authors wish to thank the staff of the Histopathology Laboratory, Ahmadu Bello University, Zaria, Nigeria, for their assistance.
Conflict of interest
The authors declare no conflict of interest.
Authors’ consent for publication
All the authors gave their consent for publication.
Ethical approval
All applicable national and institutional guidelines for the care and use of animals were followed. All procedures performed in the studied animals were following the ethical standards of the Ahmadu Bello University Committee on Animal Use and Care where the study was conducted.
Authors’ contributions
JAA and SBO participated in the study design, examination of the gross and histopathological changes, drafted the data, analyzed the data, snapping of the gross and microscopic lesions, and drafted the manuscript. LS participated in the study design and helped draft the manuscript. OO, SJE, and JOE participated in the examination of gross and histopathological changes, snapping of organs gross lesions, and helped draft the manuscript. GB, MOE, OJ, and JIS participated in the postmortem examination and helped with the snapping of gross lesions.
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