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Effect of immunosuppression on pathogenesis of peste des petits ruminants (PPR) virus infection in goats
Microbial Pathogenesis 52 (2012) 217e226
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Microbial Pathogenesis journal homepage: www.elsevier.com/locate/micpath
Effect of immunosuppression on pathogenesis of peste des petits ruminants (PPR) virus infection in goats Swapnil Pandurang Jagtap a,1, Kaushal Kishor Rajak b,1, Umesh Kumar Garg a, Arnab Sen b, Veerakyathappa Bhanuprakash b, Shashi Bhusan Sudhakar b, Vinayagamurthy Balamurugan b, Arun Patel b, Anuj Ahuja b, Raj Kumar Singh b, *, Pothukuchi Rama Vanamayya b a b
Dept. of Veterinary Pathology, College of Veterinary Science and A.H., Mhow 453446, Madhya Pradesh, India Division of Virology, Indian Veterinary Research Institute (IVRI), Mukteswar 263138, Nainital (Distt.), Uttarakhand, India
a r t i c l e i n f o
a b s t r a c t
Article history: Received 22 December 2010 Accepted 3 January 2012 Available online 10 January 2012
In this study an attempt to address the effects of immunosuppression on pathogenesis of peste des petits ruminants (PPR) virus infection was undertaken. Cyclophosphamide and dexamethasone were used to immunosuppress the animals. The drug treated animals exhibited severe leukopaenia and lymphopaenia; one of the indicators of immunosuppression. Experimental peste des petits ruminants virus (PPRV) infection was then given to both drug-induced immunosuppressed and non-immunosuppressed goats and observed their effects. Findings indicated that, the immunosuppressed goats had a short period of viremia, more extensive and severe disease advancement and higher mortality rate than the nonimmunosuppressed goats. PPRV antigen distribution in both ante-mortem and post-mortem materials was extensive and diffused in immunosuppressed animals than that of non-immunosuppressed. Some of the atypical organ(s)/tissues like liver, kidney, heart etc showed more antigen load than nonimmunosuppressed group. Histopathological and immunohistochemical studies of tissues from the two groups showed that pathological changes in the non-immunosuppressed animals were confined only to gastrointestinal tract, whereas in the immunosuppressed animals histopathological changes and PPRV antigen distribution were more extensive and diffused. The present study indicated that immunosuppression increased the extent and severity of the pathological lesions associated with peste des petits ruminants virus infection. Ó 2012 Elsevier Ltd. All rights reserved.
Keywords: Peste des petits ruminants (PPR) Immunosuppression Immunosuppressants Goat ELISA Histopathology Immunohistochemistry
1. Introduction Peste des petits ruminants (PPR) is a highly contagious and infectious viral disease of domestic and wild small ruminants that induce high morbidity and mortality [1,2]. The disease is similar to Rinderpest (RP) of cattle and characterized by severe oral necrosis, enteritis and pneumonia [3]. Sheep and goats are at present the only known natural hosts of the disease. Although cattle and pigs develop serum neutralizing antibody following experimental PPR virus infection; are not susceptible to clinical disease [4]. PPR is now enzootic in India [5,6]. It was first described in Cote d’Ivorie in West Africa [7], whereas in India in Villupuram district of Tamil Nadu [8]. The etiological agent, PPR virus has been classified under genus
* Corresponding author. Tel.: þ91 5942 286346x48; fax: þ91 5942 286347. E-mail address: [email protected] (R.K. Singh). 1 Equal contribution. 0882-4010/$ e see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.micpath.2012.01.003
Morbillivirus in the family Paramyxoviridae along with other members including rinderpest virus (RPV), measles virus (MV), canine distemper virus (CDV) and morbilliviruses of marine mammals [9,10]. Since its first report in 1942, the geographical distribution of PPR has steadily expanded to cover large regions in Africa, the Middle East and Asia [11]. PPR is considered as one of the main constraints in improving the productivity of small ruminants in the regions where it is enzootic [12]. Though PPR is a recently emerging fatal transboundary viral disease, causing huge economic loss in the Indian subcontinent [13], meagre work has been done on the effect of virus on immune status of animals. The infection leads to immunosuppression of the host [14] and results in fatal secondary bacterial infections, therefore understanding the mechanism of immunosuppression is of paramount importance. The present investigation hence was undertaken to study preliminary effects of immunosuppression on peste des petits ruminants (PPR) virus infection in goats.
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2. Results 2.1. General observations The animals that were inoculated with the virulent virus displayed a rise in temperature, characteristic of PPR, from the day onwards. The trend in the thermal response differed in between the groups. The initiation of the thermal response was similar in both the groups. The animals started showing a thermal response on day 2 following infection. However, the decline in thermal response was more rapid on 4th dpi in the immunosuppressed group vis-àvis the non-immunosuppressed group. Progressive disease induced mortality was observed in the immunosuppressed group from the 6th dpi while mortality in the non-immunosuppressed group was occurred only after 10th dpi (Fig. 1). 2.2. Drug-induced immunosuppression All animals in Group-B were subjected to drug-induced immunosuppression. The extent of immunosuppression was monitored by total white blood cell (WBC) and lymphocyte counts. 2.2.1. Total leucocyte count (TLC) after drug administration Total leucocyte counts (TLC) was monitored in Group-B and C after drug administration. In Group-B (immunosuppressed) leucopenia was observed right from day 3rd post administration of drugs (Fig. 2a) and was severe on day 5. Virus was inoculated on the 6th day to animals after ascertaining immunosuppression. The mean values of TLC in the immunosuppressed group (Group-B) were found to be significant (P < 0.05) as compared to control group (Group-C). 2.2.2. Lymphocyte counts after drug administration There was a marked lymphopenia observed in Group-B animals from day 2nd to 4th after administration of drugs (Fig. 2b). From the 4th day post administration, the lymphocyte counts showed increments and were similar to the control group on the 6th day post drug administration. The virus was administered on the 6th day post drug administration. In the non-immunosuppressed group there was no evidence of any decline in the lymphocyte count. The mean values of lymphocyte in the immunosuppressed group (Group-B) were found to be significant (P < 0.05) as compared to control group (Group-C). 2.3. Immunosuppression after PPR virus infection 2.3.1. Total leukocyte count (TLC) after virus inoculation Total leucocyte counts (TLC) was monitored in all the three groups after virus infection. Severe leucopenia was seen in Group-A
on day 4th post infection and till the animals survived, whereas in Group-B a transient rise in leukocyte count was seen from 2nd dpi, however, the level was below the normal. The mean values of TLC in Group-B were found to be significant (P < 0.05) as compared to control group (Fig. 2a). 2.3.2. Lymphocyte count after virus inoculation PPR virus is lymphotropic in nature like other morbilliviruses, therefore only lymphocyte counts (among the DLC) were monitored in the present study. Lymphocyte count was monitored in all the three groups (A, B and C). After virus infection severe lymphopenia was noticed in both the groups (A and B) from 4th dpi or till the animals survived. In Group-B there was slight rise in lymphocyte count after virus infection as compared to Group-A. The lymphocyte count in Group-B was relatively more on day 2 and 6 dpi. However, the overall pattern of lymphopaenia was similar to Group-A till day 6th dpi. From day 6 to 8, the Group-B animals showed a marginally higher lymphocyte count compared to Group-A. Group-C animals exhibited normal lymphocyte counts during the entire experimental period (Fig. 2b). The mean values of lymphocyte in all treatment groups were significant (P < 0.05) when compared with control and are as Group-A, Group-B and control Group-C. 2.4. Clinical manifestations and mortality In Group-A, all the goats exhibited typical PPR symptoms such as pyrexia (Fig. 1), mucopurulent nasal and ocular discharges, coughing, necrotic lesions in oral and nasal mucosae and bran-like deposition on tongue from 6th dpi onwards. Diarrhoea was observed in most of the goats from day 7th onwards. In Group-B (immunosuppressed), disease was severe, as the onset of clinical signs was noticed earlier as compared to the non-immunosuppressed group. Coughing and shivering started from day 3rd onwards; rectal temperature increased from 104 to 107 F. The episode of thermal reaction was less prolonged as compared to Group-A animals (Fig. 1). There was a decline in rectal temperature from day 5 onwards. Respiratory distress, rectal hemorrhages, fetid and watery diarrhoea, nasal and ocular discharge and necrosis of oral mucosa was observed from day 4th onwards. In Group-B, symptoms like huddling together, perambulatory behaviour, grunting sounds were also observed. Mortality in Group-B was recorded earlier (6e9 dpi) than the Group-A (10 dpi). Before death, animals of both groups exhibited sub normal rectal temperature, which was more pronounced in immunosuppressed animals when compared to the non-immunosuppressed ones. Group-C animals (Healthy control) didn’t show any abnormality and remained healthy throughout the experiment. Day-to-day survival rate of the animals of different groups is depicted in Fig. 3. Comparison of different parameters of signs and symptoms of PPR disease is described in Table 1. 2.5. Humoral immune response to PPRV Pattern of development of PPR specific antibody was studied in all the three groups (A, B and C). This was carried out using indigenously developed monoclonal antibody (MAb) based cELISA. Findings indicated that there was no detectable antibody response by c-ELISA test in all the groups (data not shown). 2.6. Detection of PPR viral antigen in ante and post-mortem samples
Fig. 1. Thermal reaction following virulent PPR virus infection in nonimmunosuppressed (Group-A), Immunosuppressed (Group-B) and Healthy control (Group-C). The values represent arithmetic mean in each group.
MAb based s-ELISA was employed to detect PPRV antigen from oral, nasal and rectal swabs and post-mortem tissues/organs.
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Fig. 2. a. Total Leucocyte count (TLC) in non-immunosuppressed (Group-A), Immunosuppressed (Group-B) and Healthy control (Group-C). The values represent arithmetic mean in each group. b: Lymphocyte count in non-immunosuppressed (Group-A), Immunosuppressed (Group-B) and Healthy control (Group-C).The values represent arithmetic mean in each group.
2.6.1. Swab samples In Group-A (non-immunosuppressed), PPR viral antigen was detected in nasal, oral, ocular swab on 6th dpi, whereas viral antigen in rectal swab was detected right from day 5th post infection. The virus shedding was noticed in all the swab samples till the animals survived (Fig. 4aed). In Group-B animals, early detection of antigen was noticed in ocular swab (on 3rd dpi); followed by nasal, rectal and oral on 4th dpi (Fig. 4aed). The animals in Group-C did not show any PPRV antigen in swabs. The pattern of viral antigen shedding in both the groups was almost similar; however the antigen load was more in immunosuppressed animals (Group-B) than non-immunosuppressed. Among all the swabs (oral, ocular, rectal and nasal), early detection of antigen was observed in ocular swabs in Group-B animals. In Group-A animals, early virus shedding was noticed in rectal swab. 2.6.2. Tissue samples Goats that died were subjected to post-mortem in order to ascertain the PPR specific lesions in different target organs. Different tissue/organ samples such as tongue, lungs, spleen, mesenteric lymph node, abomasum, small intestine, caecum, rectum, liver, kidney and heart were collected separately for detection of PPRV antigen. All the tissues were triturated in chilled phosphate buffer saline (PBS, pH 7.2) separately and subjected to MAb based s-ELISA to detect the presence of PPR viral antigen. PPR viral antigen load
was fairly more in immunosuppressed animals (Group-B) than nonimmunosuppressed (Group-A) barring few tissues (tongue, abomasum, caecum and rectum). Interestingly, liver showed almost equal antigen load in both the groups (Fig. 5). All the tissues of Group-C were found to be negative for PPR virus antigen. 2.7. Gross pathology The overall pathological lesions were more severe and intense in immunosuppressed (Group-B) animals than the nonimmunosuppressed one (Group-A). The details of gross pathology are summarized in Table 2. Deposition of white bran-like necrotic material on tongue was noticed in both the groups. Oral mucosa was congested in almost all cases with the presence of minor erosions on lips and tongue. The abomasal folds become congested and hemorrhagic and was more severe in Group-A. Distal parts of the small intestine were mild to moderately congested in both the groups. Lesions in the large intestine were more severe than in small intestine. The caecum showed hemorrhages, edema and thickening of corrugated mucosa in non-immunosuppressed goat. The “Zebra stripes” of congestion and haemorrhages in caecum was noticed in immunosuppressed goat. Rectum exhibited focal hemorrhages with catarrhal inflammation of mucosa in nonimmunosuppressed goat; whereas mild hemorrhages and sloughing of mucosa observed in immunosuppressed one. Mesenteric lymph nodes were swollen and oedematous in almost both groups. The spleen of affected animals exhibited enlargement. Lungs of non-immunosuppressed group showed consolidation and petechial hemorrhages in right apical and diaphragmatic lobes. Lungs of immunosuppressed group showed severe consolidation and congestion. The nasal mucosa were congested and the trachea contained blood-tinged frothy exudate. Kidney was enlarged, fragile and showed marked congestion in Group-B than Group-A. 2.8. Histopathology and immunohistochemistry
Fig. 3. Survival rate (%) of cyclophosphamide and dexamethasone-treated (Group-B) and untreated (Group-A) goats after infection with Peste des Petits ruminants virus.
Histopathological alterations were noticed in various organs such as tongue, lungs, heart, liver, kidney, abomasum, small intestine, caecum, rectum and brain in goats of different groups. The scoring of viral antigen distribution in different organs at various intervals of two groups is presented in Table 3. The overall presence of viral antigen was noticed more in case of Group-B animals than the Group-A. Lungs from all the goats showed varying degrees of histological changes involving bronchi, bronchioles and alveoli. In nonimmunosuppressed goat, lungs showed alveolar emphysema; dilated and ruptured alveoli. Presence of multinucleated giant
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Table 1 Clinical signs of PPRV infected animals. Sl. No.
Clinical signs
Groups
1
Dullness/Depression
2
Huddling together
3
Hair coat (Raised)
4
Respiratory problems (Coughing/Dyspnoea)
5
Appetite/Anorexia
6
Nasal discharge
7
Ocular discharge
8
Fever/subnormal
9
Erosions on gums
10
Diarrhoea
11
Death
A B A B A B A B A B A B A B A B A B A B A B
Days post infection 0
1
2
3
4
5
6
7
8
9
* * * * * * * * * * * * * * * * * * * *
* þ * þ * * * * * * * * * * * * * * * *
* þ * þ * * * * * * * * * * * þ * * * *
þ þþ * þþ * þ * þ * * * * * * þ þ * * * þ
þ þþ * þþ * þþ * þþ * þ þ þ þ þ þþ þþ * þ * þþ
þ þþ þ þþ þ þþ * þþ þ þþ þ þþ þ þþ þþþ þþþ þ þ * þþ
þ þþ þ þþ þ þþþ þ þþþ þþ þþ þþ þþþ þþ þþþ þþþ þþþ þ þ þ þþþ
þþ þþ þþ þþ * þþþ þþ þþþ þþ þþ þþ þþþ þþ þþþ þþþ e þ þ þþþ þþþ
þþþ þþþ þþ þþ * þþþ þþ þþþ þþþ þþþ þ þþþ þ þþþ þ e þ þ þþþ þþþ
þþþ
Δ
Δ
Δ
10
þþ * þ þþþ þ þ þ þþþ ΔΔΔΔ Δ
e: Sub normal temperature; *: Normal; þ: Mild; þþ: Moderate; þþþ: Severe; Δ: No. of goats died.
cell/syncytia was observed in lung parenchyma. Lungs of immunosuppressed group showed congested alveoli filled with inflammatory exudates predominantly with polymorphonuclear neutrophil (PMN), lymphocytes, homogenous pink coloured serous exudates and infiltration of plasma cells. Alveoli of lungs showed positive immunohistochemical reaction in non-immunosuppressed goat as evidenced by the presence of brown coloured spots. The mesenteric
lymph node (MLN) showed varying degree of lymphoid cell depletion particularly in the cortical lymphoid follicles. Lymphoid follicles exhibited mild depletion of lymphocyte in non-immunosuppressed group (Fig. 6). Lymphocytes with fragmented nuclei (apoptotic cells) were also seen in few cases in non-immunosuppressed group. Spleen of immunosuppressed group showed lymphoid depletion. There was a severe depletion of lymphocyte in spleen as well as in
Fig. 4. aed: Detection of PPR viral antigen in different swab samples by s-ELISA.
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Fig. 5. Group wise comparison of PPR viral antigen load in different tissue by monoclonal antibody based s-ELISA.
MLN in immunosuppressed goat. On day 5th post infection in immunosuppressed group, Spleen showed positive IHC reaction (Fig. 6) and in medulla, cortex and germinal centers in nonimmunosuppressed group. Mesenteric lymph node showed positive IHC reaction in lymphocyte. Sinuses, cortex and medulla of lymphoid follicles of mesenteric lymph node showed strong positive IHC reaction. Tongue exhibited mild congestion of capillaries in the stratum basale which extended up to stratum granulosum. Hydropic degeneration was observed in both the group. Ulceration and infiltration of
inflammatory cells were noticed in immunosuppressed group. Tongue showed positive IHC reaction in epithelial cells of nonimmunosuppressed group on day 10th post infection. In abomasum, there was vacuolar and hydropic degeneration and infiltration of leukocytes. Mucosal epithelium of abomasum revealed positive IHC reaction in non-immunosuppressed group. Weak positive IHC reaction noticed in lamina propria of immunosuppressed goat. Intestinal lamina propria revealed mild to severe infiltration with mononuclear cells including plasma cells and a moderate to severe vascular congestion. Necrobiotic changes were noticed in non-
Table 2 Day-wise scoring card of gross lesion in different groups infected with PPR virus.
Table 3 Detection of PPR viral antigen in various tissues by immunohistochemistry.
Sl. No
Gross lesions
Groups
Days post infection 6
1
2 3 4 5 6 7 8 9 10
Deposition of white bran-like material on tongue Consolidation of lobes of Lungs Hemorrhages in Tracheal rings Enlarged and oedematous mesenteric lymph node Enlarged Spleen Erosions and ulceration in Abomasum Streaks of congestion in Caecum Streaks of congestion in Rectum Congestion in Kidney Intestinal and mesenteric congestion
A B
7
8
9
þþ
þ
þ
Sl. No
Organs
Groups
1
Tongue
A B A B A B A B A B A B A B A B A B A B A B
10
þþ 2
A B A B A B A B A B A B A B A B A B
Lungs
þ þ
þþþ
þþþ
þ
3
Spleen
þþ
þ
þ
4
Mesentric lymph node
þþ þþþ
þþþ
þþþ
þ
5
Small Intestine
þ þ
þþ
þþ
þ
6
Abomasum
þ þ
þ
þþ
7
Caecum
þþ þþþ
þþ
þþþ
þþ
8
Rectum
þþ þþ
þþþ
þþþ
þ
9
Liver
þ þ
þþ
þþ
10
Kidney
þþ þþþ
þþ
þþþ
þþ
: No lesions; þ: Mild; þþ: Moderate; þþþ: Severe; Gr.A: Group-A - Nonimmunosuppressed; Gr.B: Group-B - Immunosuppressed; DPI: Days post infection.
11
Heart
Days post infection 6
7
8
9
þþ
þþ
þþ
þþ
þþ
þþ
þþ
þþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþ
þþ
þþ
þþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
10 þ þþ þþþ þþþ þþþ þþþ þþþ þþþ þ þ þ
Score of antigenic load: þ: Mild; þþ: Moderate; þþþ: Intense.
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Fig. 6. Histopathology and immunohistochemistry (IHC) of typical organs. A. Extensive hemorrhages and edema in mesenteric lymph node in non-immunosuppressed goat (H&EX200); B. Lymphoid follicles showing severe depletion of lymphocyte in mesenteric lymph node of immunosuppressed goats (H&EX400); C. Mesenteric lymph node showing positive IHC reaction in lymphocyte in non-immunosuppressed group (AEC substrate Mayer’s haematoxylin counterstainX400); D. Sinuses, cortex and medulla of lymphoid follicles of mesenteric lymph node showing strong positive IHC reaction in immunosuppressed group (AEC substrate Mayer’s haematoxylin counterstainX400); E. Rectum showing hemorrhages in immunosuppressed goat (H&EX400); F. Rectum showing strong positive IHC reaction in mucosal folds and villi in immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX 200).
immunosuppressed group. Villi were reduced in height and Payer’s patches showed area of lymphoid depletion in immunosuppressed group. Small intestine showed positive IHC reaction in glandular epithelium and villi of non-immunosuppressed group, whereas crypt of Liberkuhn showed positive IHC reaction in immunosuppressed group. The large intestine showed congestion of blood vessels in the mucosa and sub mucosa. Heavy infiltration of inflammatory cells and
atrophy of glandular epithelium noticed in immunosuppressed group. Rectum showed hemorrhages in immunosuppressed group along with infiltration of PMN cells. Rectum showed strong positive IHC reaction in mucosal folds and villi in non-immunosuppressed group (Fig. 6). In the liver, hepatocytes were swollen having pale and finely granular cytoplasm. Infiltration of PMN cells were noticed in non-
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immunosuppressed group. In immunosuppressed group, liver showed less infiltration of inflammatory cells. Swollen hepatocyte and reduced sinusoidal space in immunosuppressed group was also seen. Nucleus of hepatocytes exhibited strong positive IHC reaction in both the group (Fig. 7). In kidneys, the epithelial cells lining the proximal convoluted tubules were swollen. Swollen glomeruli, infiltration of inflammatory cells and degenerative changes in the tubules of kidneys of non-immunosuppressed was noticed. Severe atrophic changes were also observed in non-immunosuppressed group; whereas, mild glomerular atrophy was noticed in immunosuppressed group. Infiltration of inflammatory cells in interstitial tubules was seen in immunosuppressed group. Fatty changes and hemorrhages were noticed in kidney of immunosuppressed group. The tubular epithelium of kidney of both the group showed positive IHC reaction (Fig. 7). Interstitial connective tissue showed positive IHC reaction in non-immunosuppressed group. Heart showed congestion, degenerative changes and focal area of necrosis in nonimmunosuppressed group. In case of immunosuppressed group, severe hemorrhages were noticed after infection. Cardiomyocytes of heart showed positive IHC reaction in immunosuppressed group. Brain did not show any significant change except mild congestion. 3. Discussion and conclusion The results of this study clearly show that cyclophosphamide and dexamethasone induced immunosuppression enhances PPRV infection. Cyclophosphamide and dexamethasone affect both humoral and cell mediated immune responses [15] and both components of the immune response are important in controlling the infection [16,17]. The treated goats in Group-B did not elicit humoral immune response; however, pathological changes were extensive and severe, the onset of mortality was earlier than the untreated infected goats in Group-A. Similar changes have been observed with West Nile virus experimentally infected animals’ and immunosuppressed with alkylating agents, radiation, corticosteroids, and thymectomy [18]. Thus, we believe that the immunosuppressed goat model offers some insight into the pathogenesis of PPRV infection in immunosuppressed goats and why in a flock some goats are at higher risk to PPR followed by death, when they become infected with PPRV. PPR virus causes a severe disease in its natural host i.e. goat and sheep, associated with high morbidity and mortality. PPR virus is a lymphotropic virus similar to Rinderpest [19]. To study the effect of immunosuppression of PPR virus infected goats (natural host), animals were classified in to three groups; viz. non-immunosuppressed (Group-A), immunosuppressed (Group-B) and healthy control (Group-C). Experiment was designed to study the effects of PPR virus both in immunosuppressed and nonimmunosuppressed animals. Parameters such as TLC, DLC (lymphocyte), antibody response against PPR virus and viral antigen detection in different clinical samples of PPR infected animals were undertaken. In order to induce immunosuppression, animals were injected with cyclophosphamide on the first day along with tapering doses of dexamethasone as per the schedule described by Koptopoulos et al. [20]. Cyclophosphamide is a potent immunosuppressive drug especially at higher dose. It is toxic for resting and dividing cells, especially for dividing immunocompetent cells. It impairs both B and T cell response, especially primary immune response. It tends to destroy more B cells than T cells. It also suppresses macrophage function and has anti-inflammatory effects. It has a half-life of about 6 h and largely excreted through the kidney [21]. On the other hand, dexamethasone is a corticosteroid. Corticosteroid blocks all NF-kb mediated processes including cytokine synthesis. Administration or excessive release of corticosteroids in vivo
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typically induces leukocytosis, neutrophilia, lymphopenia, eosinopenia, basopenia and monocytosis or monocytopenia depending upon the species involved [21,22]. Initially, total leukocyte count (TLC) was monitored in Group-B and C following drug administration. There was a severe fall in total leukocyte count from day 2nd onwards which remained up to day 5th post drug administration. In case of DLC, there was a marked lymphopenia observed in Group-B animals from day 2nd to 4th after administration of drugs, implying the immunosuppression. Then it reached normal on the day of virus infection. There was no change in TLC and DLC of healthy control. The results obtained in the present study are in agreement with the study conducted elsewhere [20]. As expected, the needle passaged PPR challenge virus (PPRV, Izatnagar/94) induced typical PPR specific symptoms which has been reconfirmed on several occasions [23,24] including this study. The severity of clinical symptoms was more in immunosuppressed group than the non-immunosuppressed. In the present study, the animals treated with immunosuppressant drugs revealed immunosuppression as indicated by haematology (TLC and Lymphocyte). Hence the immunosuppressed animals manifested severe symptoms than the animals infected without immunosuppression. Humoral immunity is an important arm of immune system. Morbillivirus in general are supposed to induce strong humoral antibody response after vaccination or infection. After inoculation of virulent virus in both the groups, the blood was tested in MAb based c-ELISA for the detection of specific antibody against PPR virus. It is interesting to note that, as the samples were available only up to day 8th post infection, there was no immune response against PPR virus. Similarly, Rajak et al. [14] failed to detect PPR specific antibodies during the 1st week in PPR infected goats. They also noticed that the challenge virus interferes with the generation of immune response against other antigen/s, suggestive of a marked but transient immunosuppression caused by the virus during the initial phase of infection. In the present study, immunosuppressed goats, died on or before day 9th post infection and non-immunosuppressed on day 10th. Therefore, immune system of affected goats could not produce detectable amount of PPR specific antibody by this time. This was generally due to the fact that the animals showed mortality prior to the minimum window required 21 days for seroconversion. Transmission of PPR infection is by close contact between infected and susceptible animals. To diagnose and confirm the infection of goats by PPR virus as early as possible, nasal, ocular, oral and rectal swabs were tested by s-ELISA. These are the classical samples for the identification of early virus shedding. These samples were collected from all the three groups on different days. It is interesting to note that, there was an early detection of PPR virus from ocular swab in immunosuppressed group on day 3rd onwards; whereas detection of PPR antigen was observed in rectal swab on day 5th onwards of non-immunosuppressed. Nasal and oral swabs were found to be positive in both the groups. These findings corroborate with the earlier observations [25,26]. Excretions of PPR virus through these routes are responsible for rapid transmission of PPRV from sick to in-contact susceptible animals during epizootics. Goats died on different days were conducted post-mortem in order to examine for PPR specific lesions. Different tissue samples such as tongue, lungs, spleen, mesenteric lymph node, abomasum, small intestine, caecum, rectum, liver, kidney and heart from these goats were collected separately. All the tissues then subjected to MAb based s-ELISA to detect the presence of PPR viral antigen, histopathology and immunohistochemistry. As expected, tissues of immunosuppressed animals exhibited more viral antigen load (as exhibited by optical density values in s-ELISA) than the normally
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Fig. 7. Histopathology and immunohistochemistry (IHC) of atypical organs. A. Liver showing infiltration of PMN cells in non-immunosuppressed goats (H&EX400); B. Haemorrhages and swollen cells observed in liver cells with variable degree of necrobiotic changes in immunosuppressed goat (H&EX400); C. Nucleus of hepatocytes of liver showing strong positive IHC reaction in non-immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX400.); D. Nucleus of hepatocytes of liver showing strong positive IHC reaction in immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX400); E. Swollen glomeruli, Infiltration of inflammatory cells and degenerative changes in tubules in kidney of non-immunosuppressed goat (H&EX400); F. Kidney showing fatty changes and hemorrhages in immunosuppressed goat (H&EX400); G. The Tubular epithelium of kidney showing positive IHC reaction in non-immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX1000); H. The Tubular epithelium of kidney showing strong positive IHC reaction in immunosuppressed group (AEC substrate Mayer’s haematoxylin counterstain.X1000).
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infected one. Interestingly, some of the intestinal tissues (abomasums, caecum and rectum) and tongue epithelium showed more antigen load in normally infected animals (Group-A) than the immunosuppressed one. This could be due to less opportunity of virus to infect the intestinal tissues in immunosuppressed animals, as they died earlier than the normal one. Another reason could be the early diarrhoea in immunosuppressed animals, which might have leached out the antigen from intestine; however, we have not tested the faeces. In the immunosuppressed group, the antigen processing might have been slower leading to a low viral localization. As the extrusion of the virus depends on efficient virus localization, immunosuppression might have an adverse effect on virus localization. It remains to be seen that which effect of the immune response has correlation with the maturation of the virus. However in this study a “cause and effect” phenomenon has been speculated between a robust immune response and complete virus maturation/replication in vivo. The histopathology and immunohistochemistry of most of the target tissues (intestines, tongue, lungs and lymph nodes) showed positive signal for PPR viral antigen which is in agreement with Kumar et al. [27]. The lesions were more extensive in immunosuppressed groups. Further, some of the atypical organs like liver, kidney and heart also exhibited positive reaction in immunosuppressed animals. It is to hypothesise here that the diffused nature of PPR viral infection in immunosuppressed animals is probably the result of decreased immune response and persistent viremia which leads to the involvement of atypical organs. The results of this study clearly show that cyclophosphamide and dexamethasone induced marked lymphopenia observed in Group-B animals after administration of drugs, which is an indicator of immunosuppression. The immunosuppression further precipitated the PPRV infection. Cyclophosphamide and dexamethasone affect both the humoral and cell mediated immune responses, and both components of the immune response are important in evading the infection. The treated goats in Group-B had no humoral immune response, more extensive and severe disease progression, and an earlier mortality than the untreated infected goats in Group-A. The same phenomenon has been observed with West Nile virus in experimentally infected animals immunosuppressed with alkylating agents, radiation, corticosteroids, and thymectomy. Thus, we believe that the immunosuppressed goat model offers some insight into the pathogenesis of PPRV infection in immunosuppressed goats.
The goats were randomly divided into two groups (Group-A: Nonimmunosuppressed; Group-B: Immunosuppressed) consisting of four goats each. Both these groups were infected with the virus and one control group (Group: C) comprising two goats were neither infected nor immunosuppressed. Of the two infected groups, Group-B was immunosuppressed with cyclophosphamide and dexamethasone [20]. Each animal in the infected groups (Group-A and -B) was inoculated with virulent PPRV (PPRV, Izatnagar/94) using 10% spleenic tissue suspension by subcutaneous route. A total of 5 ml (ml) tissue suspension was given to each animal in two equal divided doses on each axilla. The goats of the control group were similarly administered with 5 ml of sterile normal saline solution (NSS) as a placebo. Animals of different groups were maintained in separate animal facility throughout the experiment with no cross handling. All the animals were monitored diurnally for temperature reaction and PPR specific disease symptoms up to 10 days or till animal survived.
4. Materials and methods
All the ante-mortem (swabs) and post-mortem (tissues/organs) samples were subjected to a monoclonal antibody based sandwichELISA test as described by Singh et al. [29] in order to detect the PPR virus. The test uses a polyclonal antibody as capture antibody and monoclonal antibody to ‘N’ protein of PPRV as detection antibody.
4.1. PPR virus A highly virulent peste des petits ruminants virus (PPRV, Izatnagar/94) isolated from an outbreak in Uttar Pradesh, India in 1994 is being maintained by animal-to-animal passages at National Morbillivirus Referral Laboratory, Indian Veterinary Research Institute, Mukteswar was used [23]. Affected animals were sacrificed and the spleen and mesenteric lymph nodes were harvested and stored at 80 C for subsequent passage/challenge. The challenge virus was used as a 10% spleenic suspension that was injected subcutaneously in volumes of 5 ml in an adult hilly goat with an average body weight of 15e20 kgs.
4.3. Drug-induced immunosuppression All four animals of Group-B were subjected to drug-induced immunosuppression. For immunosuppression, both cyclophosphamide and dexamethasone injection was used. The dose and schedule of immunisation are shown in Table 4 [20]. The level of immunosuppression was monitored by total white blood cell (WBC) and lymphocyte counts [22]. 4.4. Haematological studies Heparinized (20 IU/ml) blood samples in sterile vaccutainer tubes were collected a day before the experimental infection and on alternate days following infection to determine total leukocyte count (TLC) and lymphocyte count (DLC) [22]. 4.5. Detection of antibodies to PPR virus using competitive-ELISA Detection of PPRV antibodies in the sera samples collected from PPR infected and control animals was carried out using monoclonal antibody (directed to H protein) based competitive-ELISA (c-ELISA) test [27]. Samples that had a percent inhibition (PI) value of >40% were considered positive and samples having a PI value of <40% were considered negative. 4.6. Detection of PPR virus antigen using sandwich-ELISA
Table 4 The dose and schedule of immunisation in different groups of goats. Sl. No Group
No of Treatment animals
1
Group-A 04
2
Group-B 04
3
Group-C 02
4.2. Animal experimental design Apparently 10 healthy hill goats of either sex aged 12 months were ascertained for their PPR free status by a monoclonal antibody based competitive-ELISA [28] and were used in the present study.
Infection with 10% spleenic tissue suspension of PPR virulent virus Injection of Dexamethasone on 5 consecutive days [For the first 3 days @ 4 mg/kg body wt, on 4th day @ 3 mg/kg body wt and on the 5th day @ 2.5 mg/kg body wt (i/v)]. Injection of cyclophosphamide on 1st day @ 25 mg/kg body wt. (i/v) Infection with 10% spleenic tissue suspension of PPR virulent virus on 6th day. Normal saline solution (NSS)
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4.7. Histopathology The histopathological procedure was followed as per the method described by Luna [30] with little modification. Briefly, after fixation in 10% buffered neutral formalin solution for 24e48 h, small pieces of tissues were cut and kept under tap water overnight for washing. The tissues were then dehydrated in ascending grades of alcohol, cleared in cedar wood oil and embedded in paraffin wax (60e62 C melting point). Sections of 4e6 micron thickness were cut through a Spencer’s rotary microtome and stained with Haematoxylin and Eosin. 4.8. Immunohistochemistry The test was carried out using ExtrAvidin Peroxidase staining kit, Mouse (SigmaeAldrich St Louis, USA) in order to detect PPRV antigen from tissue samples as per the instruction manual. Briefly, tissue sections were digested with protease (0.1%). The endogenous peroxidase activity was quenched with 3% hydrogen peroxide (H2O2). Non-specific sites were blocked by incubating each section with 3% bovine serum albumin (BSA) and the sections were covered with 1:20 diluted primary antibody (Monoclonal antibody to N protein of PPRV). An aminoethylcarbazole (AEC) chromogen substrate system (SigmaeAldrich St Louis, USA) was applied for colour reaction. Peste des petits ruminants virus e positive mesenteric lymph node tissues, previously confirmed with RT-PCR, were used as positive controls. Lung sections from two goats (1 year old) with no clinicopathological evidences of PPRV infection were used as negative control. Acknowledgement The authors are grateful to the Head of the Division of Virology and the Director of the Institute for providing the necessary facilities to carry out this work. References [1] Furley CW, Taylor WP, Obi TU. An outbreak of peste des petits ruminants in a zoological collection. Vet Rec 1987;121:443e7. [2] Almeida RS, Keita D, Libeau G, Albina E. Control of ruminant morbillivirus replication by small interfering RNA. J Gen Virol 2007;88:2307e11. [3] Nduaka O, Ihemelandu EC. Observation of pneumoenteritis complex in dwarf goats in Eastern states of Nigeria. Bull Epizoot Dis Afr 1973;21:87. [4] Anderson J, McKay JA. The detection of antibodies against peste des petits ruminants virus in cattle, sheep and goats and the possible implication to rinderpest control programme. Epidemiol Infect 1994;112:225e31. [5] Nanda YP, Chatterjee AA, Purohit K, Diallo A, Innui K, Sharma RN, et al. The isolation of peste des petits ruminants virus from Northern India. Vet Microbiol 1996;51:207e16. [6] Shaila MS, Peter AB, Varalakshmi P, Apte M, Rajendran MP, Anbumani SP. Peste des petits ruminants in Tamilnadu goats. Indian Vet J 1996;73:587e8.
[7] Gargadennec L, Lalanne A. La peste des petits ruminants. Bull Serv Zoot Epiz A O F 1942;5(1):16e21. [8] Shaila MS, Purushothaman V, Bhavasar D, Venugopal K, Venkatesan RA. Peste des petits ruminants of sheep in India. Vet Rec 1989;125:602. [9] Barrett T, Visser IKJ, Mamaev L, Goatley L, van Bressem MF, Osterhaus DME. Dolphin and porpoise morbilli viruses are genetically distinct from phocine distemper. Virol 1993;193:1010e2. [10] Van Regenmortel MHV, Fauquet CM, Bishop DHL. Virus taxonomy. Seventh report of the international committee on taxonomy of viruses. San Diego: Academic Press; 2000. [11] http://www.fao.org/ag/againfo/programmes/en/empres/news_201010.html. [12] Stem C. An economic analysis of the prevention of peste des petits ruminants in Nigerian goats. Prev Vet Med 1999;16:141e50. [13] Venkataramanan R, Bandyopadhyay SK, Oberoi MS. Present status and strategies for the control of transboundary and other economically important animal diseases in India: a review. Ind J Anim Sci 2005;75:456e64. [14] Rajak KK, Sreenivasa BP, Hosamani M, Singh RP, Singh SK, Singh RK, et al. Experimental studies on immunosuppressive effects of Peste des petits ruminants (PPR) virus in goats. Comp Immunol Microbiol Infect Dis 2005;28: 287e96. [15] Ahmed AR, Hombal SM. Cyclophosphamide (Cytoxan*). J Am Acad Dermatol 1984;11:1115e26. [16] Chambers TJ, Diamond MS. Pathogenesis of flavivirus encephalitis. Adv Virus Res 2003;60:273e342. [17] Wang T, Fikrig E. Immunity to West Nile virus. Curr Opin Immunol; 2004. [18] Mateo R, Xiao SY, Guzman H, Lei H, Amelia PA, Rosa TD, et al. Effects of immunosuppression on West Nile virus infection in Hamsters. Am J Trop Med Hyg 2006;75(2). [19] Fraser CM. The Merck Vet. Mann. 6th ed. USA: Merck and Co. Inc.; 1986. [20] Koptopoulos G, Papanastasopoulou M, Lekkas S, Skaragas G, Papadopoulos O. Immunosuppression in goats by dexamethasone and cyclophosphamide. Comp Immunol Microbiol Infect Dis 1992;15(4):235e42. [21] Tizzard IR. Veterinary Immunology. An introduction. Vth ed.; 1996 [pp. 457e458]. [22] Jain NC. Schalm’s Veterinary Haematology. 4th ed. Philadelphia: Lea and Febiges; 1986 [pp. 121e123]. [23] Sreenivasa BP, Dhar P, Singh RP, Bandopadhyay SK. Development of peste des petits ruminants (PPR) challenge virus from a field isolate. Presented at “XIV annual conference and national seminar on management of viral diseases with emphasis on global trade and WTO regime” of Indian Virological Society, January 18e20, 2002, Indian veterinary research institute, Hebbal Bangalore, Karnataka-560024. [24] Singh RP. Production and characterization of monoclonal antibodies to peste des petits ruminants (PPR) virus. Ph.D. thesis. Submitted to IVRI, Deemed University, Izatnagar, India 2002. [25] Abegunde AA, Adu FD. Excretion of the virus of peste des petits ruminants by goats. Bull Anim Health Prod Afr 1977;25(3):307e11. [26] Gibbs EPT, Taylor WP, Lawman MP, Bryant J. Classification of the peste des petits ruminants virus as the fourth member of the genus Morbillivirus. Intervirol 1979;11:268e74. [27] Kumar P, Tripathi BN, Sharma AK, Kumar R, Sreenivasa BP, Singh RP, et al. Pathological and immunohistochemical study of experimental peste des petits ruminants virus infection in goats. J Vet Med 2004;51:153e9. [28] Singh RP, Sreenivasa BP, Dhar P, Bandyopadhyay SK. A sandwich-ELISA for the diagnosis of Peste des petits ruminants (PPR) infection in small ruminants using anti-nucleocapsid protein monoclonal antibody. Arch Virol 2004b;149: 2155e70. [29] Singh RP, Sreenivasa BP, Dhar P, Shah LC, Bandyopadhyay SK. Development of Monoclonal antibody based competitive- ELISA for detection and titration of antibodies to peste des petits ruminants virus. Vet Microbiol 2004a;98:3e15. [30] Luna LG. Manual on histologic staining methods of the armed force institute of pathology. 3rd ed. USA: McGraw-Hill Book Company; 1968 [pp. 32e46].
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Microbial Pathogenesis journal homepage: www.elsevier.com/locate/micpath
Effect of immunosuppression on pathogenesis of peste des petits ruminants (PPR) virus infection in goats Swapnil Pandurang Jagtap a,1, Kaushal Kishor Rajak b,1, Umesh Kumar Garg a, Arnab Sen b, Veerakyathappa Bhanuprakash b, Shashi Bhusan Sudhakar b, Vinayagamurthy Balamurugan b, Arun Patel b, Anuj Ahuja b, Raj Kumar Singh b, *, Pothukuchi Rama Vanamayya b a b
Dept. of Veterinary Pathology, College of Veterinary Science and A.H., Mhow 453446, Madhya Pradesh, India Division of Virology, Indian Veterinary Research Institute (IVRI), Mukteswar 263138, Nainital (Distt.), Uttarakhand, India
a r t i c l e i n f o
a b s t r a c t
Article history: Received 22 December 2010 Accepted 3 January 2012 Available online 10 January 2012
In this study an attempt to address the effects of immunosuppression on pathogenesis of peste des petits ruminants (PPR) virus infection was undertaken. Cyclophosphamide and dexamethasone were used to immunosuppress the animals. The drug treated animals exhibited severe leukopaenia and lymphopaenia; one of the indicators of immunosuppression. Experimental peste des petits ruminants virus (PPRV) infection was then given to both drug-induced immunosuppressed and non-immunosuppressed goats and observed their effects. Findings indicated that, the immunosuppressed goats had a short period of viremia, more extensive and severe disease advancement and higher mortality rate than the nonimmunosuppressed goats. PPRV antigen distribution in both ante-mortem and post-mortem materials was extensive and diffused in immunosuppressed animals than that of non-immunosuppressed. Some of the atypical organ(s)/tissues like liver, kidney, heart etc showed more antigen load than nonimmunosuppressed group. Histopathological and immunohistochemical studies of tissues from the two groups showed that pathological changes in the non-immunosuppressed animals were confined only to gastrointestinal tract, whereas in the immunosuppressed animals histopathological changes and PPRV antigen distribution were more extensive and diffused. The present study indicated that immunosuppression increased the extent and severity of the pathological lesions associated with peste des petits ruminants virus infection. Ó 2012 Elsevier Ltd. All rights reserved.
Keywords: Peste des petits ruminants (PPR) Immunosuppression Immunosuppressants Goat ELISA Histopathology Immunohistochemistry
1. Introduction Peste des petits ruminants (PPR) is a highly contagious and infectious viral disease of domestic and wild small ruminants that induce high morbidity and mortality [1,2]. The disease is similar to Rinderpest (RP) of cattle and characterized by severe oral necrosis, enteritis and pneumonia [3]. Sheep and goats are at present the only known natural hosts of the disease. Although cattle and pigs develop serum neutralizing antibody following experimental PPR virus infection; are not susceptible to clinical disease [4]. PPR is now enzootic in India [5,6]. It was first described in Cote d’Ivorie in West Africa [7], whereas in India in Villupuram district of Tamil Nadu [8]. The etiological agent, PPR virus has been classified under genus
* Corresponding author. Tel.: þ91 5942 286346x48; fax: þ91 5942 286347. E-mail address: [email protected] (R.K. Singh). 1 Equal contribution. 0882-4010/$ e see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.micpath.2012.01.003
Morbillivirus in the family Paramyxoviridae along with other members including rinderpest virus (RPV), measles virus (MV), canine distemper virus (CDV) and morbilliviruses of marine mammals [9,10]. Since its first report in 1942, the geographical distribution of PPR has steadily expanded to cover large regions in Africa, the Middle East and Asia [11]. PPR is considered as one of the main constraints in improving the productivity of small ruminants in the regions where it is enzootic [12]. Though PPR is a recently emerging fatal transboundary viral disease, causing huge economic loss in the Indian subcontinent [13], meagre work has been done on the effect of virus on immune status of animals. The infection leads to immunosuppression of the host [14] and results in fatal secondary bacterial infections, therefore understanding the mechanism of immunosuppression is of paramount importance. The present investigation hence was undertaken to study preliminary effects of immunosuppression on peste des petits ruminants (PPR) virus infection in goats.
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2. Results 2.1. General observations The animals that were inoculated with the virulent virus displayed a rise in temperature, characteristic of PPR, from the day onwards. The trend in the thermal response differed in between the groups. The initiation of the thermal response was similar in both the groups. The animals started showing a thermal response on day 2 following infection. However, the decline in thermal response was more rapid on 4th dpi in the immunosuppressed group vis-àvis the non-immunosuppressed group. Progressive disease induced mortality was observed in the immunosuppressed group from the 6th dpi while mortality in the non-immunosuppressed group was occurred only after 10th dpi (Fig. 1). 2.2. Drug-induced immunosuppression All animals in Group-B were subjected to drug-induced immunosuppression. The extent of immunosuppression was monitored by total white blood cell (WBC) and lymphocyte counts. 2.2.1. Total leucocyte count (TLC) after drug administration Total leucocyte counts (TLC) was monitored in Group-B and C after drug administration. In Group-B (immunosuppressed) leucopenia was observed right from day 3rd post administration of drugs (Fig. 2a) and was severe on day 5. Virus was inoculated on the 6th day to animals after ascertaining immunosuppression. The mean values of TLC in the immunosuppressed group (Group-B) were found to be significant (P < 0.05) as compared to control group (Group-C). 2.2.2. Lymphocyte counts after drug administration There was a marked lymphopenia observed in Group-B animals from day 2nd to 4th after administration of drugs (Fig. 2b). From the 4th day post administration, the lymphocyte counts showed increments and were similar to the control group on the 6th day post drug administration. The virus was administered on the 6th day post drug administration. In the non-immunosuppressed group there was no evidence of any decline in the lymphocyte count. The mean values of lymphocyte in the immunosuppressed group (Group-B) were found to be significant (P < 0.05) as compared to control group (Group-C). 2.3. Immunosuppression after PPR virus infection 2.3.1. Total leukocyte count (TLC) after virus inoculation Total leucocyte counts (TLC) was monitored in all the three groups after virus infection. Severe leucopenia was seen in Group-A
on day 4th post infection and till the animals survived, whereas in Group-B a transient rise in leukocyte count was seen from 2nd dpi, however, the level was below the normal. The mean values of TLC in Group-B were found to be significant (P < 0.05) as compared to control group (Fig. 2a). 2.3.2. Lymphocyte count after virus inoculation PPR virus is lymphotropic in nature like other morbilliviruses, therefore only lymphocyte counts (among the DLC) were monitored in the present study. Lymphocyte count was monitored in all the three groups (A, B and C). After virus infection severe lymphopenia was noticed in both the groups (A and B) from 4th dpi or till the animals survived. In Group-B there was slight rise in lymphocyte count after virus infection as compared to Group-A. The lymphocyte count in Group-B was relatively more on day 2 and 6 dpi. However, the overall pattern of lymphopaenia was similar to Group-A till day 6th dpi. From day 6 to 8, the Group-B animals showed a marginally higher lymphocyte count compared to Group-A. Group-C animals exhibited normal lymphocyte counts during the entire experimental period (Fig. 2b). The mean values of lymphocyte in all treatment groups were significant (P < 0.05) when compared with control and are as Group-A, Group-B and control Group-C. 2.4. Clinical manifestations and mortality In Group-A, all the goats exhibited typical PPR symptoms such as pyrexia (Fig. 1), mucopurulent nasal and ocular discharges, coughing, necrotic lesions in oral and nasal mucosae and bran-like deposition on tongue from 6th dpi onwards. Diarrhoea was observed in most of the goats from day 7th onwards. In Group-B (immunosuppressed), disease was severe, as the onset of clinical signs was noticed earlier as compared to the non-immunosuppressed group. Coughing and shivering started from day 3rd onwards; rectal temperature increased from 104 to 107 F. The episode of thermal reaction was less prolonged as compared to Group-A animals (Fig. 1). There was a decline in rectal temperature from day 5 onwards. Respiratory distress, rectal hemorrhages, fetid and watery diarrhoea, nasal and ocular discharge and necrosis of oral mucosa was observed from day 4th onwards. In Group-B, symptoms like huddling together, perambulatory behaviour, grunting sounds were also observed. Mortality in Group-B was recorded earlier (6e9 dpi) than the Group-A (10 dpi). Before death, animals of both groups exhibited sub normal rectal temperature, which was more pronounced in immunosuppressed animals when compared to the non-immunosuppressed ones. Group-C animals (Healthy control) didn’t show any abnormality and remained healthy throughout the experiment. Day-to-day survival rate of the animals of different groups is depicted in Fig. 3. Comparison of different parameters of signs and symptoms of PPR disease is described in Table 1. 2.5. Humoral immune response to PPRV Pattern of development of PPR specific antibody was studied in all the three groups (A, B and C). This was carried out using indigenously developed monoclonal antibody (MAb) based cELISA. Findings indicated that there was no detectable antibody response by c-ELISA test in all the groups (data not shown). 2.6. Detection of PPR viral antigen in ante and post-mortem samples
Fig. 1. Thermal reaction following virulent PPR virus infection in nonimmunosuppressed (Group-A), Immunosuppressed (Group-B) and Healthy control (Group-C). The values represent arithmetic mean in each group.
MAb based s-ELISA was employed to detect PPRV antigen from oral, nasal and rectal swabs and post-mortem tissues/organs.
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Fig. 2. a. Total Leucocyte count (TLC) in non-immunosuppressed (Group-A), Immunosuppressed (Group-B) and Healthy control (Group-C). The values represent arithmetic mean in each group. b: Lymphocyte count in non-immunosuppressed (Group-A), Immunosuppressed (Group-B) and Healthy control (Group-C).The values represent arithmetic mean in each group.
2.6.1. Swab samples In Group-A (non-immunosuppressed), PPR viral antigen was detected in nasal, oral, ocular swab on 6th dpi, whereas viral antigen in rectal swab was detected right from day 5th post infection. The virus shedding was noticed in all the swab samples till the animals survived (Fig. 4aed). In Group-B animals, early detection of antigen was noticed in ocular swab (on 3rd dpi); followed by nasal, rectal and oral on 4th dpi (Fig. 4aed). The animals in Group-C did not show any PPRV antigen in swabs. The pattern of viral antigen shedding in both the groups was almost similar; however the antigen load was more in immunosuppressed animals (Group-B) than non-immunosuppressed. Among all the swabs (oral, ocular, rectal and nasal), early detection of antigen was observed in ocular swabs in Group-B animals. In Group-A animals, early virus shedding was noticed in rectal swab. 2.6.2. Tissue samples Goats that died were subjected to post-mortem in order to ascertain the PPR specific lesions in different target organs. Different tissue/organ samples such as tongue, lungs, spleen, mesenteric lymph node, abomasum, small intestine, caecum, rectum, liver, kidney and heart were collected separately for detection of PPRV antigen. All the tissues were triturated in chilled phosphate buffer saline (PBS, pH 7.2) separately and subjected to MAb based s-ELISA to detect the presence of PPR viral antigen. PPR viral antigen load
was fairly more in immunosuppressed animals (Group-B) than nonimmunosuppressed (Group-A) barring few tissues (tongue, abomasum, caecum and rectum). Interestingly, liver showed almost equal antigen load in both the groups (Fig. 5). All the tissues of Group-C were found to be negative for PPR virus antigen. 2.7. Gross pathology The overall pathological lesions were more severe and intense in immunosuppressed (Group-B) animals than the nonimmunosuppressed one (Group-A). The details of gross pathology are summarized in Table 2. Deposition of white bran-like necrotic material on tongue was noticed in both the groups. Oral mucosa was congested in almost all cases with the presence of minor erosions on lips and tongue. The abomasal folds become congested and hemorrhagic and was more severe in Group-A. Distal parts of the small intestine were mild to moderately congested in both the groups. Lesions in the large intestine were more severe than in small intestine. The caecum showed hemorrhages, edema and thickening of corrugated mucosa in non-immunosuppressed goat. The “Zebra stripes” of congestion and haemorrhages in caecum was noticed in immunosuppressed goat. Rectum exhibited focal hemorrhages with catarrhal inflammation of mucosa in nonimmunosuppressed goat; whereas mild hemorrhages and sloughing of mucosa observed in immunosuppressed one. Mesenteric lymph nodes were swollen and oedematous in almost both groups. The spleen of affected animals exhibited enlargement. Lungs of non-immunosuppressed group showed consolidation and petechial hemorrhages in right apical and diaphragmatic lobes. Lungs of immunosuppressed group showed severe consolidation and congestion. The nasal mucosa were congested and the trachea contained blood-tinged frothy exudate. Kidney was enlarged, fragile and showed marked congestion in Group-B than Group-A. 2.8. Histopathology and immunohistochemistry
Fig. 3. Survival rate (%) of cyclophosphamide and dexamethasone-treated (Group-B) and untreated (Group-A) goats after infection with Peste des Petits ruminants virus.
Histopathological alterations were noticed in various organs such as tongue, lungs, heart, liver, kidney, abomasum, small intestine, caecum, rectum and brain in goats of different groups. The scoring of viral antigen distribution in different organs at various intervals of two groups is presented in Table 3. The overall presence of viral antigen was noticed more in case of Group-B animals than the Group-A. Lungs from all the goats showed varying degrees of histological changes involving bronchi, bronchioles and alveoli. In nonimmunosuppressed goat, lungs showed alveolar emphysema; dilated and ruptured alveoli. Presence of multinucleated giant
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Table 1 Clinical signs of PPRV infected animals. Sl. No.
Clinical signs
Groups
1
Dullness/Depression
2
Huddling together
3
Hair coat (Raised)
4
Respiratory problems (Coughing/Dyspnoea)
5
Appetite/Anorexia
6
Nasal discharge
7
Ocular discharge
8
Fever/subnormal
9
Erosions on gums
10
Diarrhoea
11
Death
A B A B A B A B A B A B A B A B A B A B A B
Days post infection 0
1
2
3
4
5
6
7
8
9
* * * * * * * * * * * * * * * * * * * *
* þ * þ * * * * * * * * * * * * * * * *
* þ * þ * * * * * * * * * * * þ * * * *
þ þþ * þþ * þ * þ * * * * * * þ þ * * * þ
þ þþ * þþ * þþ * þþ * þ þ þ þ þ þþ þþ * þ * þþ
þ þþ þ þþ þ þþ * þþ þ þþ þ þþ þ þþ þþþ þþþ þ þ * þþ
þ þþ þ þþ þ þþþ þ þþþ þþ þþ þþ þþþ þþ þþþ þþþ þþþ þ þ þ þþþ
þþ þþ þþ þþ * þþþ þþ þþþ þþ þþ þþ þþþ þþ þþþ þþþ e þ þ þþþ þþþ
þþþ þþþ þþ þþ * þþþ þþ þþþ þþþ þþþ þ þþþ þ þþþ þ e þ þ þþþ þþþ
þþþ
Δ
Δ
Δ
10
þþ * þ þþþ þ þ þ þþþ ΔΔΔΔ Δ
e: Sub normal temperature; *: Normal; þ: Mild; þþ: Moderate; þþþ: Severe; Δ: No. of goats died.
cell/syncytia was observed in lung parenchyma. Lungs of immunosuppressed group showed congested alveoli filled with inflammatory exudates predominantly with polymorphonuclear neutrophil (PMN), lymphocytes, homogenous pink coloured serous exudates and infiltration of plasma cells. Alveoli of lungs showed positive immunohistochemical reaction in non-immunosuppressed goat as evidenced by the presence of brown coloured spots. The mesenteric
lymph node (MLN) showed varying degree of lymphoid cell depletion particularly in the cortical lymphoid follicles. Lymphoid follicles exhibited mild depletion of lymphocyte in non-immunosuppressed group (Fig. 6). Lymphocytes with fragmented nuclei (apoptotic cells) were also seen in few cases in non-immunosuppressed group. Spleen of immunosuppressed group showed lymphoid depletion. There was a severe depletion of lymphocyte in spleen as well as in
Fig. 4. aed: Detection of PPR viral antigen in different swab samples by s-ELISA.
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Fig. 5. Group wise comparison of PPR viral antigen load in different tissue by monoclonal antibody based s-ELISA.
MLN in immunosuppressed goat. On day 5th post infection in immunosuppressed group, Spleen showed positive IHC reaction (Fig. 6) and in medulla, cortex and germinal centers in nonimmunosuppressed group. Mesenteric lymph node showed positive IHC reaction in lymphocyte. Sinuses, cortex and medulla of lymphoid follicles of mesenteric lymph node showed strong positive IHC reaction. Tongue exhibited mild congestion of capillaries in the stratum basale which extended up to stratum granulosum. Hydropic degeneration was observed in both the group. Ulceration and infiltration of
inflammatory cells were noticed in immunosuppressed group. Tongue showed positive IHC reaction in epithelial cells of nonimmunosuppressed group on day 10th post infection. In abomasum, there was vacuolar and hydropic degeneration and infiltration of leukocytes. Mucosal epithelium of abomasum revealed positive IHC reaction in non-immunosuppressed group. Weak positive IHC reaction noticed in lamina propria of immunosuppressed goat. Intestinal lamina propria revealed mild to severe infiltration with mononuclear cells including plasma cells and a moderate to severe vascular congestion. Necrobiotic changes were noticed in non-
Table 2 Day-wise scoring card of gross lesion in different groups infected with PPR virus.
Table 3 Detection of PPR viral antigen in various tissues by immunohistochemistry.
Sl. No
Gross lesions
Groups
Days post infection 6
1
2 3 4 5 6 7 8 9 10
Deposition of white bran-like material on tongue Consolidation of lobes of Lungs Hemorrhages in Tracheal rings Enlarged and oedematous mesenteric lymph node Enlarged Spleen Erosions and ulceration in Abomasum Streaks of congestion in Caecum Streaks of congestion in Rectum Congestion in Kidney Intestinal and mesenteric congestion
A B
7
8
9
þþ
þ
þ
Sl. No
Organs
Groups
1
Tongue
A B A B A B A B A B A B A B A B A B A B A B
10
þþ 2
A B A B A B A B A B A B A B A B A B
Lungs
þ þ
þþþ
þþþ
þ
3
Spleen
þþ
þ
þ
4
Mesentric lymph node
þþ þþþ
þþþ
þþþ
þ
5
Small Intestine
þ þ
þþ
þþ
þ
6
Abomasum
þ þ
þ
þþ
7
Caecum
þþ þþþ
þþ
þþþ
þþ
8
Rectum
þþ þþ
þþþ
þþþ
þ
9
Liver
þ þ
þþ
þþ
10
Kidney
þþ þþþ
þþ
þþþ
þþ
: No lesions; þ: Mild; þþ: Moderate; þþþ: Severe; Gr.A: Group-A - Nonimmunosuppressed; Gr.B: Group-B - Immunosuppressed; DPI: Days post infection.
11
Heart
Days post infection 6
7
8
9
þþ
þþ
þþ
þþ
þþ
þþ
þþ
þþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþ
þþ
þþ
þþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þþþ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
10 þ þþ þþþ þþþ þþþ þþþ þþþ þþþ þ þ þ
Score of antigenic load: þ: Mild; þþ: Moderate; þþþ: Intense.
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Fig. 6. Histopathology and immunohistochemistry (IHC) of typical organs. A. Extensive hemorrhages and edema in mesenteric lymph node in non-immunosuppressed goat (H&EX200); B. Lymphoid follicles showing severe depletion of lymphocyte in mesenteric lymph node of immunosuppressed goats (H&EX400); C. Mesenteric lymph node showing positive IHC reaction in lymphocyte in non-immunosuppressed group (AEC substrate Mayer’s haematoxylin counterstainX400); D. Sinuses, cortex and medulla of lymphoid follicles of mesenteric lymph node showing strong positive IHC reaction in immunosuppressed group (AEC substrate Mayer’s haematoxylin counterstainX400); E. Rectum showing hemorrhages in immunosuppressed goat (H&EX400); F. Rectum showing strong positive IHC reaction in mucosal folds and villi in immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX 200).
immunosuppressed group. Villi were reduced in height and Payer’s patches showed area of lymphoid depletion in immunosuppressed group. Small intestine showed positive IHC reaction in glandular epithelium and villi of non-immunosuppressed group, whereas crypt of Liberkuhn showed positive IHC reaction in immunosuppressed group. The large intestine showed congestion of blood vessels in the mucosa and sub mucosa. Heavy infiltration of inflammatory cells and
atrophy of glandular epithelium noticed in immunosuppressed group. Rectum showed hemorrhages in immunosuppressed group along with infiltration of PMN cells. Rectum showed strong positive IHC reaction in mucosal folds and villi in non-immunosuppressed group (Fig. 6). In the liver, hepatocytes were swollen having pale and finely granular cytoplasm. Infiltration of PMN cells were noticed in non-
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immunosuppressed group. In immunosuppressed group, liver showed less infiltration of inflammatory cells. Swollen hepatocyte and reduced sinusoidal space in immunosuppressed group was also seen. Nucleus of hepatocytes exhibited strong positive IHC reaction in both the group (Fig. 7). In kidneys, the epithelial cells lining the proximal convoluted tubules were swollen. Swollen glomeruli, infiltration of inflammatory cells and degenerative changes in the tubules of kidneys of non-immunosuppressed was noticed. Severe atrophic changes were also observed in non-immunosuppressed group; whereas, mild glomerular atrophy was noticed in immunosuppressed group. Infiltration of inflammatory cells in interstitial tubules was seen in immunosuppressed group. Fatty changes and hemorrhages were noticed in kidney of immunosuppressed group. The tubular epithelium of kidney of both the group showed positive IHC reaction (Fig. 7). Interstitial connective tissue showed positive IHC reaction in non-immunosuppressed group. Heart showed congestion, degenerative changes and focal area of necrosis in nonimmunosuppressed group. In case of immunosuppressed group, severe hemorrhages were noticed after infection. Cardiomyocytes of heart showed positive IHC reaction in immunosuppressed group. Brain did not show any significant change except mild congestion. 3. Discussion and conclusion The results of this study clearly show that cyclophosphamide and dexamethasone induced immunosuppression enhances PPRV infection. Cyclophosphamide and dexamethasone affect both humoral and cell mediated immune responses [15] and both components of the immune response are important in controlling the infection [16,17]. The treated goats in Group-B did not elicit humoral immune response; however, pathological changes were extensive and severe, the onset of mortality was earlier than the untreated infected goats in Group-A. Similar changes have been observed with West Nile virus experimentally infected animals’ and immunosuppressed with alkylating agents, radiation, corticosteroids, and thymectomy [18]. Thus, we believe that the immunosuppressed goat model offers some insight into the pathogenesis of PPRV infection in immunosuppressed goats and why in a flock some goats are at higher risk to PPR followed by death, when they become infected with PPRV. PPR virus causes a severe disease in its natural host i.e. goat and sheep, associated with high morbidity and mortality. PPR virus is a lymphotropic virus similar to Rinderpest [19]. To study the effect of immunosuppression of PPR virus infected goats (natural host), animals were classified in to three groups; viz. non-immunosuppressed (Group-A), immunosuppressed (Group-B) and healthy control (Group-C). Experiment was designed to study the effects of PPR virus both in immunosuppressed and nonimmunosuppressed animals. Parameters such as TLC, DLC (lymphocyte), antibody response against PPR virus and viral antigen detection in different clinical samples of PPR infected animals were undertaken. In order to induce immunosuppression, animals were injected with cyclophosphamide on the first day along with tapering doses of dexamethasone as per the schedule described by Koptopoulos et al. [20]. Cyclophosphamide is a potent immunosuppressive drug especially at higher dose. It is toxic for resting and dividing cells, especially for dividing immunocompetent cells. It impairs both B and T cell response, especially primary immune response. It tends to destroy more B cells than T cells. It also suppresses macrophage function and has anti-inflammatory effects. It has a half-life of about 6 h and largely excreted through the kidney [21]. On the other hand, dexamethasone is a corticosteroid. Corticosteroid blocks all NF-kb mediated processes including cytokine synthesis. Administration or excessive release of corticosteroids in vivo
223
typically induces leukocytosis, neutrophilia, lymphopenia, eosinopenia, basopenia and monocytosis or monocytopenia depending upon the species involved [21,22]. Initially, total leukocyte count (TLC) was monitored in Group-B and C following drug administration. There was a severe fall in total leukocyte count from day 2nd onwards which remained up to day 5th post drug administration. In case of DLC, there was a marked lymphopenia observed in Group-B animals from day 2nd to 4th after administration of drugs, implying the immunosuppression. Then it reached normal on the day of virus infection. There was no change in TLC and DLC of healthy control. The results obtained in the present study are in agreement with the study conducted elsewhere [20]. As expected, the needle passaged PPR challenge virus (PPRV, Izatnagar/94) induced typical PPR specific symptoms which has been reconfirmed on several occasions [23,24] including this study. The severity of clinical symptoms was more in immunosuppressed group than the non-immunosuppressed. In the present study, the animals treated with immunosuppressant drugs revealed immunosuppression as indicated by haematology (TLC and Lymphocyte). Hence the immunosuppressed animals manifested severe symptoms than the animals infected without immunosuppression. Humoral immunity is an important arm of immune system. Morbillivirus in general are supposed to induce strong humoral antibody response after vaccination or infection. After inoculation of virulent virus in both the groups, the blood was tested in MAb based c-ELISA for the detection of specific antibody against PPR virus. It is interesting to note that, as the samples were available only up to day 8th post infection, there was no immune response against PPR virus. Similarly, Rajak et al. [14] failed to detect PPR specific antibodies during the 1st week in PPR infected goats. They also noticed that the challenge virus interferes with the generation of immune response against other antigen/s, suggestive of a marked but transient immunosuppression caused by the virus during the initial phase of infection. In the present study, immunosuppressed goats, died on or before day 9th post infection and non-immunosuppressed on day 10th. Therefore, immune system of affected goats could not produce detectable amount of PPR specific antibody by this time. This was generally due to the fact that the animals showed mortality prior to the minimum window required 21 days for seroconversion. Transmission of PPR infection is by close contact between infected and susceptible animals. To diagnose and confirm the infection of goats by PPR virus as early as possible, nasal, ocular, oral and rectal swabs were tested by s-ELISA. These are the classical samples for the identification of early virus shedding. These samples were collected from all the three groups on different days. It is interesting to note that, there was an early detection of PPR virus from ocular swab in immunosuppressed group on day 3rd onwards; whereas detection of PPR antigen was observed in rectal swab on day 5th onwards of non-immunosuppressed. Nasal and oral swabs were found to be positive in both the groups. These findings corroborate with the earlier observations [25,26]. Excretions of PPR virus through these routes are responsible for rapid transmission of PPRV from sick to in-contact susceptible animals during epizootics. Goats died on different days were conducted post-mortem in order to examine for PPR specific lesions. Different tissue samples such as tongue, lungs, spleen, mesenteric lymph node, abomasum, small intestine, caecum, rectum, liver, kidney and heart from these goats were collected separately. All the tissues then subjected to MAb based s-ELISA to detect the presence of PPR viral antigen, histopathology and immunohistochemistry. As expected, tissues of immunosuppressed animals exhibited more viral antigen load (as exhibited by optical density values in s-ELISA) than the normally
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Fig. 7. Histopathology and immunohistochemistry (IHC) of atypical organs. A. Liver showing infiltration of PMN cells in non-immunosuppressed goats (H&EX400); B. Haemorrhages and swollen cells observed in liver cells with variable degree of necrobiotic changes in immunosuppressed goat (H&EX400); C. Nucleus of hepatocytes of liver showing strong positive IHC reaction in non-immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX400.); D. Nucleus of hepatocytes of liver showing strong positive IHC reaction in immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX400); E. Swollen glomeruli, Infiltration of inflammatory cells and degenerative changes in tubules in kidney of non-immunosuppressed goat (H&EX400); F. Kidney showing fatty changes and hemorrhages in immunosuppressed goat (H&EX400); G. The Tubular epithelium of kidney showing positive IHC reaction in non-immunosuppressed group (AEC substrate. Mayer’s haematoxylin counterstainX1000); H. The Tubular epithelium of kidney showing strong positive IHC reaction in immunosuppressed group (AEC substrate Mayer’s haematoxylin counterstain.X1000).
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infected one. Interestingly, some of the intestinal tissues (abomasums, caecum and rectum) and tongue epithelium showed more antigen load in normally infected animals (Group-A) than the immunosuppressed one. This could be due to less opportunity of virus to infect the intestinal tissues in immunosuppressed animals, as they died earlier than the normal one. Another reason could be the early diarrhoea in immunosuppressed animals, which might have leached out the antigen from intestine; however, we have not tested the faeces. In the immunosuppressed group, the antigen processing might have been slower leading to a low viral localization. As the extrusion of the virus depends on efficient virus localization, immunosuppression might have an adverse effect on virus localization. It remains to be seen that which effect of the immune response has correlation with the maturation of the virus. However in this study a “cause and effect” phenomenon has been speculated between a robust immune response and complete virus maturation/replication in vivo. The histopathology and immunohistochemistry of most of the target tissues (intestines, tongue, lungs and lymph nodes) showed positive signal for PPR viral antigen which is in agreement with Kumar et al. [27]. The lesions were more extensive in immunosuppressed groups. Further, some of the atypical organs like liver, kidney and heart also exhibited positive reaction in immunosuppressed animals. It is to hypothesise here that the diffused nature of PPR viral infection in immunosuppressed animals is probably the result of decreased immune response and persistent viremia which leads to the involvement of atypical organs. The results of this study clearly show that cyclophosphamide and dexamethasone induced marked lymphopenia observed in Group-B animals after administration of drugs, which is an indicator of immunosuppression. The immunosuppression further precipitated the PPRV infection. Cyclophosphamide and dexamethasone affect both the humoral and cell mediated immune responses, and both components of the immune response are important in evading the infection. The treated goats in Group-B had no humoral immune response, more extensive and severe disease progression, and an earlier mortality than the untreated infected goats in Group-A. The same phenomenon has been observed with West Nile virus in experimentally infected animals immunosuppressed with alkylating agents, radiation, corticosteroids, and thymectomy. Thus, we believe that the immunosuppressed goat model offers some insight into the pathogenesis of PPRV infection in immunosuppressed goats.
The goats were randomly divided into two groups (Group-A: Nonimmunosuppressed; Group-B: Immunosuppressed) consisting of four goats each. Both these groups were infected with the virus and one control group (Group: C) comprising two goats were neither infected nor immunosuppressed. Of the two infected groups, Group-B was immunosuppressed with cyclophosphamide and dexamethasone [20]. Each animal in the infected groups (Group-A and -B) was inoculated with virulent PPRV (PPRV, Izatnagar/94) using 10% spleenic tissue suspension by subcutaneous route. A total of 5 ml (ml) tissue suspension was given to each animal in two equal divided doses on each axilla. The goats of the control group were similarly administered with 5 ml of sterile normal saline solution (NSS) as a placebo. Animals of different groups were maintained in separate animal facility throughout the experiment with no cross handling. All the animals were monitored diurnally for temperature reaction and PPR specific disease symptoms up to 10 days or till animal survived.
4. Materials and methods
All the ante-mortem (swabs) and post-mortem (tissues/organs) samples were subjected to a monoclonal antibody based sandwichELISA test as described by Singh et al. [29] in order to detect the PPR virus. The test uses a polyclonal antibody as capture antibody and monoclonal antibody to ‘N’ protein of PPRV as detection antibody.
4.1. PPR virus A highly virulent peste des petits ruminants virus (PPRV, Izatnagar/94) isolated from an outbreak in Uttar Pradesh, India in 1994 is being maintained by animal-to-animal passages at National Morbillivirus Referral Laboratory, Indian Veterinary Research Institute, Mukteswar was used [23]. Affected animals were sacrificed and the spleen and mesenteric lymph nodes were harvested and stored at 80 C for subsequent passage/challenge. The challenge virus was used as a 10% spleenic suspension that was injected subcutaneously in volumes of 5 ml in an adult hilly goat with an average body weight of 15e20 kgs.
4.3. Drug-induced immunosuppression All four animals of Group-B were subjected to drug-induced immunosuppression. For immunosuppression, both cyclophosphamide and dexamethasone injection was used. The dose and schedule of immunisation are shown in Table 4 [20]. The level of immunosuppression was monitored by total white blood cell (WBC) and lymphocyte counts [22]. 4.4. Haematological studies Heparinized (20 IU/ml) blood samples in sterile vaccutainer tubes were collected a day before the experimental infection and on alternate days following infection to determine total leukocyte count (TLC) and lymphocyte count (DLC) [22]. 4.5. Detection of antibodies to PPR virus using competitive-ELISA Detection of PPRV antibodies in the sera samples collected from PPR infected and control animals was carried out using monoclonal antibody (directed to H protein) based competitive-ELISA (c-ELISA) test [27]. Samples that had a percent inhibition (PI) value of >40% were considered positive and samples having a PI value of <40% were considered negative. 4.6. Detection of PPR virus antigen using sandwich-ELISA
Table 4 The dose and schedule of immunisation in different groups of goats. Sl. No Group
No of Treatment animals
1
Group-A 04
2
Group-B 04
3
Group-C 02
4.2. Animal experimental design Apparently 10 healthy hill goats of either sex aged 12 months were ascertained for their PPR free status by a monoclonal antibody based competitive-ELISA [28] and were used in the present study.
Infection with 10% spleenic tissue suspension of PPR virulent virus Injection of Dexamethasone on 5 consecutive days [For the first 3 days @ 4 mg/kg body wt, on 4th day @ 3 mg/kg body wt and on the 5th day @ 2.5 mg/kg body wt (i/v)]. Injection of cyclophosphamide on 1st day @ 25 mg/kg body wt. (i/v) Infection with 10% spleenic tissue suspension of PPR virulent virus on 6th day. Normal saline solution (NSS)
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4.7. Histopathology The histopathological procedure was followed as per the method described by Luna [30] with little modification. Briefly, after fixation in 10% buffered neutral formalin solution for 24e48 h, small pieces of tissues were cut and kept under tap water overnight for washing. The tissues were then dehydrated in ascending grades of alcohol, cleared in cedar wood oil and embedded in paraffin wax (60e62 C melting point). Sections of 4e6 micron thickness were cut through a Spencer’s rotary microtome and stained with Haematoxylin and Eosin. 4.8. Immunohistochemistry The test was carried out using ExtrAvidin Peroxidase staining kit, Mouse (SigmaeAldrich St Louis, USA) in order to detect PPRV antigen from tissue samples as per the instruction manual. Briefly, tissue sections were digested with protease (0.1%). The endogenous peroxidase activity was quenched with 3% hydrogen peroxide (H2O2). Non-specific sites were blocked by incubating each section with 3% bovine serum albumin (BSA) and the sections were covered with 1:20 diluted primary antibody (Monoclonal antibody to N protein of PPRV). An aminoethylcarbazole (AEC) chromogen substrate system (SigmaeAldrich St Louis, USA) was applied for colour reaction. Peste des petits ruminants virus e positive mesenteric lymph node tissues, previously confirmed with RT-PCR, were used as positive controls. Lung sections from two goats (1 year old) with no clinicopathological evidences of PPRV infection were used as negative control. Acknowledgement The authors are grateful to the Head of the Division of Virology and the Director of the Institute for providing the necessary facilities to carry out this work. References [1] Furley CW, Taylor WP, Obi TU. An outbreak of peste des petits ruminants in a zoological collection. Vet Rec 1987;121:443e7. [2] Almeida RS, Keita D, Libeau G, Albina E. Control of ruminant morbillivirus replication by small interfering RNA. J Gen Virol 2007;88:2307e11. [3] Nduaka O, Ihemelandu EC. Observation of pneumoenteritis complex in dwarf goats in Eastern states of Nigeria. Bull Epizoot Dis Afr 1973;21:87. [4] Anderson J, McKay JA. The detection of antibodies against peste des petits ruminants virus in cattle, sheep and goats and the possible implication to rinderpest control programme. Epidemiol Infect 1994;112:225e31. [5] Nanda YP, Chatterjee AA, Purohit K, Diallo A, Innui K, Sharma RN, et al. The isolation of peste des petits ruminants virus from Northern India. Vet Microbiol 1996;51:207e16. [6] Shaila MS, Peter AB, Varalakshmi P, Apte M, Rajendran MP, Anbumani SP. Peste des petits ruminants in Tamilnadu goats. Indian Vet J 1996;73:587e8.
[7] Gargadennec L, Lalanne A. La peste des petits ruminants. Bull Serv Zoot Epiz A O F 1942;5(1):16e21. [8] Shaila MS, Purushothaman V, Bhavasar D, Venugopal K, Venkatesan RA. Peste des petits ruminants of sheep in India. Vet Rec 1989;125:602. [9] Barrett T, Visser IKJ, Mamaev L, Goatley L, van Bressem MF, Osterhaus DME. Dolphin and porpoise morbilli viruses are genetically distinct from phocine distemper. Virol 1993;193:1010e2. [10] Van Regenmortel MHV, Fauquet CM, Bishop DHL. Virus taxonomy. Seventh report of the international committee on taxonomy of viruses. San Diego: Academic Press; 2000. [11] http://www.fao.org/ag/againfo/programmes/en/empres/news_201010.html. [12] Stem C. An economic analysis of the prevention of peste des petits ruminants in Nigerian goats. Prev Vet Med 1999;16:141e50. [13] Venkataramanan R, Bandyopadhyay SK, Oberoi MS. Present status and strategies for the control of transboundary and other economically important animal diseases in India: a review. Ind J Anim Sci 2005;75:456e64. [14] Rajak KK, Sreenivasa BP, Hosamani M, Singh RP, Singh SK, Singh RK, et al. Experimental studies on immunosuppressive effects of Peste des petits ruminants (PPR) virus in goats. Comp Immunol Microbiol Infect Dis 2005;28: 287e96. [15] Ahmed AR, Hombal SM. Cyclophosphamide (Cytoxan*). J Am Acad Dermatol 1984;11:1115e26. [16] Chambers TJ, Diamond MS. Pathogenesis of flavivirus encephalitis. Adv Virus Res 2003;60:273e342. [17] Wang T, Fikrig E. Immunity to West Nile virus. Curr Opin Immunol; 2004. [18] Mateo R, Xiao SY, Guzman H, Lei H, Amelia PA, Rosa TD, et al. Effects of immunosuppression on West Nile virus infection in Hamsters. Am J Trop Med Hyg 2006;75(2). [19] Fraser CM. The Merck Vet. Mann. 6th ed. USA: Merck and Co. Inc.; 1986. [20] Koptopoulos G, Papanastasopoulou M, Lekkas S, Skaragas G, Papadopoulos O. Immunosuppression in goats by dexamethasone and cyclophosphamide. Comp Immunol Microbiol Infect Dis 1992;15(4):235e42. [21] Tizzard IR. Veterinary Immunology. An introduction. Vth ed.; 1996 [pp. 457e458]. [22] Jain NC. Schalm’s Veterinary Haematology. 4th ed. Philadelphia: Lea and Febiges; 1986 [pp. 121e123]. [23] Sreenivasa BP, Dhar P, Singh RP, Bandopadhyay SK. Development of peste des petits ruminants (PPR) challenge virus from a field isolate. Presented at “XIV annual conference and national seminar on management of viral diseases with emphasis on global trade and WTO regime” of Indian Virological Society, January 18e20, 2002, Indian veterinary research institute, Hebbal Bangalore, Karnataka-560024. [24] Singh RP. Production and characterization of monoclonal antibodies to peste des petits ruminants (PPR) virus. Ph.D. thesis. Submitted to IVRI, Deemed University, Izatnagar, India 2002. [25] Abegunde AA, Adu FD. Excretion of the virus of peste des petits ruminants by goats. Bull Anim Health Prod Afr 1977;25(3):307e11. [26] Gibbs EPT, Taylor WP, Lawman MP, Bryant J. Classification of the peste des petits ruminants virus as the fourth member of the genus Morbillivirus. Intervirol 1979;11:268e74. [27] Kumar P, Tripathi BN, Sharma AK, Kumar R, Sreenivasa BP, Singh RP, et al. Pathological and immunohistochemical study of experimental peste des petits ruminants virus infection in goats. J Vet Med 2004;51:153e9. [28] Singh RP, Sreenivasa BP, Dhar P, Bandyopadhyay SK. A sandwich-ELISA for the diagnosis of Peste des petits ruminants (PPR) infection in small ruminants using anti-nucleocapsid protein monoclonal antibody. Arch Virol 2004b;149: 2155e70. [29] Singh RP, Sreenivasa BP, Dhar P, Shah LC, Bandyopadhyay SK. Development of Monoclonal antibody based competitive- ELISA for detection and titration of antibodies to peste des petits ruminants virus. Vet Microbiol 2004a;98:3e15. [30] Luna LG. Manual on histologic staining methods of the armed force institute of pathology. 3rd ed. USA: McGraw-Hill Book Company; 1968 [pp. 32e46].