Detailed coverage of invasive species threatening livelihoods and the environment worldwide
Generate a print friendly version containing only the sections you need.
Generate reportRotaviruses are the commonest cause of severe, acute viral gastroenteritis in man, mammals and birds worldwide, with most of cases occurring in young during the winter. In developing countries, rotaviruses have been estimated to cause 600,000 to 870,000 deaths each year, accounting for about 20-25% of all deaths due to diarrhoea and 6% of all deaths among children (DeZoysa and Feachem, 1985; Institute of Medicine, 1986). Though occasionally implicated as a foodborne illnesses, the virus is mainly transmitted through the faecal-oral route and is only excreted only the faeces. Prevalence studies show that 29.4% of infected animals excrete the virus in faeces. However, other routes of transmission, such as waterborne or airborne (respiratory) routes, have also been suggested. There is evidence that zoonotic transmission of rotaviruses, or at least rotavirus genes, can occur. The rapid movement of people of diverse populations throughout the world allows the introduction and spread of rotaviruses generated in developing countries through zoonotic transmission and ressortment. The incubation period of rotavirus diarrhoeal illness has been estimated to be less than 48 h.
The rotaviral infection, which mostly occurs as concurrent infection with other enteropathogens, leads to diarrhoea within 2-4 days of consumption of contaminated food/water and lasts for 2-5 days. Mortality in affected persons can be as high as 20%. Rotaviral diarrhoea in neonatal animals causes major economic loss directly through mortality and therapy and indirectly from poor growth after clinical disease. The factors which influence rotavirus infection and its clinical severity include the age of the animal, immune status of the dam and absorption of colostral antibody, ambient temperature, degree of viral exposure, the occurrence of weaning, and the presence of the other enteropathogens (Saif and Smith, 1985; Bridger and Pocock, 1986).
Rotaviruses derived the name from Latin word, ‘rota’, meaning ‘wheel’, because of their morphology. Rotaviruses are members of the Reoviridae family and, possess a genome consisting of 11 segments of double-stranded RNA (dsRNA) enclosed in a triple-layered capsid. These have been classified into 7 groups (A-G) and 4 serotypes. Groups A, B, and C have been found in both humans and animals while groups D, E, F and G have been found to infect not only humans, but also birds like chickens, turkeys and mammals like calves, piglets, foals, dogs, cats, deer, rabbits and mice. Viruses of human origin share a common antigen with Nebraska calf diarrhoea virus (NCDV), simian agent (SA)-11 and offal agent of sheep and goat. Most of the rotaviruses have been found to cross inter-species barriers under experimental conditions. Natural transmission of the virus has been documented between cattle and pigs species; and also between man and calves. The two outer capsid proteins, VP4 and VP7, the main antigenic determinants, independently elicit neutralizing antibodies, and induce protective immunity (Estes, 2001). Based on either antigenic or genetic characterization, 15 VP7 gene alleles (each corresponding to a G serotype) and 23 VP4 gene alleles (each corresponding to P genotypes) have been recognized. Due to the lack of appropriate antibody reagents, out of the 23 P genotypes identified so far, only 14 P serotypes and 3 subtypes have been established (Estes, 2001;Hoshino et al.,2002; Martella et al.,2003; Liprandi et al., 2003).
The clinical signs of rotavirus gastroenteritis are not distinct enough to permit a specific diagnosis. Laboratory diagnosis of rotavirus infections requires identifying the virus in faeces or rectal swab specimens or demonstrating a fourfold or greater increase in antibody to a rotavirus antigen between acute- and convalescent-phase sera. Samples of intestinal mucosa from several sections of the small and large intestine should be submitted chilled for virus detection and possible isolation. Electron microscopic examination of faecal material has remained a standard diagnostic technique. The other methods to detect rotavirus in stool and rectal swab specimens include electron microscopy, radio-immunoassay, counterimmunoelectro-osmophoresis, centrifuging of clinical material onto tissue culture cells followed by immunofluorescence, inoculation of tissue cultures, latex agglutination, reverse passive haemagglutination assay, polyacrylamide gel electrophoresis, dot hybridization, polymerase chain reaction, and enzyme-linked immunosorbent assay (ELISA).
Antibodies to a variety of rotavirus antigens develop during infection; neutralizing antibodies to both major neutralizing proteins, VP4 and VP7, have been shown to induce protection in animal studies.
No specific therapy is advocated to deal with rotaviral diarrhoea. Anti-microbial agents are used both orally and parenterally to treat the possible co-infection with enteric and systemic bacterial infections. The primary aim of treatment of rotavirus enteritis is the replacement, by intravenous or oral routes, of fluids and electrolytes lost by diarrhoea. Rotavirus vaccine development has focused on the delivery of live attenuated rotavirus strains by the oral route. The initial Jennerian approach involving bovine (RIT4237, WC3) or rhesus rotavirus (RRV) vaccine candidates has shown that these vaccines are safe, well tolerated, and immunogenic.
| Animal name | Context | Life stage | System |
|---|---|---|---|
| Bos indicus (zebu) | Domesticated host | Cattle and Buffaloes|All Stages | |
| Bos taurus (cattle) | Domesticated host | Cattle and Buffaloes|All Stages | |
| Bovidae | Wild host | ||
| Bubalus bubalis (Asian water buffalo) | Cattle and Buffaloes|All Stages | ||
| Camelus bactrianus (Bactrian camel) | Domesticated host | ||
| Camelus dromedarius (dromedary camel) | Domesticated host | ||
| Canis familiaris (dogs) | Domesticated host | ||
| Capra hircus (goats) | Domesticated host | Sheep and Goats|Lamb | |
| Equus caballus (horses) | Domesticated host | ||
| Felis catus (cat) | Domesticated host | ||
| Gallus gallus domesticus (chickens) | Domesticated host | Poultry|All Stages | |
| Lama glama (llamas) | Wild host | ||
| Mus musculus (house mouse) | Experimental settings | ||
| Oryctolagus cuniculus (rabbits) | Domesticated host; Wild host | ||
| Ovis aries (sheep) | Domesticated host | Sheep and Goats|Lamb | |
| Phasianus (pheasants) | Wild host | ||
| Primates | Wild host | ||
| Sus scrofa (pigs) | Pigs|All Stages |
Humans of all ages are susceptible to rotavirus infection. Children of 6 months to 2 years of age, premature infants, the elderly, and the immunocompromised are particularly prone to more severe symptoms caused by infection with Group A rotavirus. Calves of both sexes are affected. In India, calves of crossbred or exotic breeds were more susceptible to rotavirus infection than indigenous animals (Chauhan and Singh, 1996). Clinical illness was associated with dams shedding virus and with low night temperatures (Bulgin et al., 1989). The factors which influence rotavirus infection and its clinical severity include the age of the animal, immune status of the dam and absorption of colostral antibody, ambient temperature, degree of viral exposure, the occurrence of weaning, and the presence of the other enteropathogens (Saif and Smith, 1985; Bridger and Pocock, 1986). The mortality is highest in the youngest animals, which have received insufficient colostrums and are subjected to severe weather conditions. Colostral serum antibody in newborn animals does not protect animals against clinical disease (Snodgrass and Wells, 1978). Calves born in crowded herds or grouped in large numbers were more likely to be positive for rotavirus. Moreover, as the number of calves in the herd increased, the risk of rotavirus infection also increased (Erdoan et al., 2003).
The prevalence of Group A rotavirus was more related to meterological changes than age of the lambs as the number of diarrhoeic lambs with rotavirus infection was found to increase in spring during which temperature and humidity ranged between 7.34 and 28.9°C and 34.28 and 82.58%, respectively (Wani et al., 2004a).
Larger herd size and younger weaning age predisposed to rotavirus infection (Dewey et al., 2003). Pigs maintained in all-in all-out nurseries were 3.4 times more likely to have a positive Group A rotavirus diagnosis than pigs in continuous flow facilities. Changes in Group A rotavirus disease herd status have been found to be associated with changes in farm management practices, including farm expansion, early weaning, and all-in all-out production (Dewey et al., 2003).
A comparatively higher percentage of serological reactions was detected in pigs reared under intensive farming systems than that in open grazing systems (Barman et al., 2003). Low relative humidity, high population density and high concentration of sows that have farrowed are some of the predisposing factors to rotavirus infection (Alfieri et al., 1999). Piglets aged 3 and 5 weeks gave the highest virus isolation rates (Alfieri et al., 1991). This pattern of infection was attributed to the non-intensive husbandry methods in the villages, with less opportunity for transmission to occur than in intensive production facilities (Alpers et al., 1991).
Rotavirus infections have a worldwide distribution and are a common cause of neonatal diarrhoea in many mammalian and avian species. Geographically, rotaviruses have been found in all places wherever they have been searched for. In temperate countries, a consistent seasonal pattern of rotavirus infections is exhibited with peaks in the winter. In epidemiological studies conducted in developing countries, rotaviruses accounted for, a median of 8% of all diarrhoeal episodes, 28% of outpatients or clinic visits for diarrhoea, and 34% of hospitalizations of young children for diarrhoea.
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
Last updated: 10 Jan 2020| Continent/Country/Region | Distribution | Last Reported | Origin | First Reported | Invasive | Reference | Notes |
|---|---|---|---|---|---|---|---|
Africa |
|||||||
| Cameroon | Present | ||||||
| Congo, Democratic Republic of the | Present, Localized | ||||||
| Ethiopia | Present, Widespread | ||||||
| Ghana | Present | ||||||
| Morocco | Present, Widespread | ||||||
| Nigeria | Present, Widespread | ||||||
| South Africa | Present, Localized | ||||||
| Sudan | Present, Widespread | ||||||
| Tunisia | Present, Widespread | ||||||
| Zimbabwe | Present, Widespread | ||||||
Asia |
|||||||
| Bahrain | Present | Native | |||||
| Bangladesh | Present, Widespread | Native | |||||
| Bhutan | Present, Widespread | ||||||
| China | Present | Present based on regional distribution. | |||||
| -Anhui | Present | ||||||
| -Fujian | Present, Localized | Original citation: Fujian Provincial Institute of Animal Husbandry an (1981) | |||||
| -Henan | Present | ||||||
| -Hunan | Present | ||||||
| -Jiangsu | Present | ||||||
| -Qinghai | Present, Localized | ||||||
| -Sichuan | Present, Localized | ||||||
| -Xinjiang | Present, Widespread | ||||||
| -Yunnan | Present, Widespread | ||||||
| India | Present | ||||||
| -Assam | Present, Widespread | ||||||
| -Chandigarh | Present, Widespread | ||||||
| -Delhi | Present, Widespread | ||||||
| -Gujarat | Present, Widespread | ||||||
| -Haryana | Present, Widespread | ||||||
| -Jammu and Kashmir | Present, Widespread | ||||||
| -Karnataka | Present, Widespread | ||||||
| -Madhya Pradesh | Present, Widespread | ||||||
| -Maharashtra | Present, Widespread | ||||||
| -Manipur | Present, Widespread | ||||||
| -Punjab | Present, Widespread | ||||||
| -Tamil Nadu | Present, Widespread | ||||||
| -Uttar Pradesh | Present, Widespread | ||||||
| -West Bengal | Present, Widespread | ||||||
| Indonesia | Present | Present based on regional distribution. | |||||
| -Java | Present | ||||||
| -Lesser Sunda Islands | Present | ||||||
| Iran | Present, Widespread | ||||||
| Iraq | Present | ||||||
| Israel | Present, Localized | ||||||
| Japan | Present | ||||||
| -Hokkaido | Present, Widespread | Native | |||||
| -Honshu | Present, Widespread | ||||||
| -Kyushu | Present, Widespread | ||||||
| -Ryukyu Islands | Present, Widespread | ||||||
| -Shikoku | Present, Widespread | ||||||
| Malaysia | Present | Present based on regional distribution. | |||||
| -Peninsular Malaysia | Present, Widespread | ||||||
| North Korea | Present | ||||||
| South Korea | Present | ||||||
| Sri Lanka | Present, Localized | ||||||
| Taiwan | Present | ||||||
| Thailand | Present, Widespread | ||||||
| United Arab Emirates | Present, Widespread | ||||||
| Uzbekistan | Present | ||||||
| Vietnam | Present | ||||||
Europe |
|||||||
| Albania | Present, Widespread | ||||||
| Austria | Present, Widespread | ||||||
| Belarus | Present | ||||||
| Belgium | Present, Widespread | ||||||
| Bulgaria | Present, Localized | ||||||
| Czechia | Present, Localized | ||||||
| Czechoslovakia | Present, Localized | ||||||
| Federal Republic of Yugoslavia | Present, Localized | ||||||
| Denmark | Present, Localized | ||||||
| Finland | Present, Localized | ||||||
| France | Present, Localized | ||||||
| Germany | Present, Localized | ||||||
| Hungary | Present, Localized | ||||||
| Ireland | Present, Localized | ||||||
| Italy | Present, Localized | ||||||
| Netherlands | Present, Localized | ||||||
| Poland | Present, Widespread | ||||||
| Portugal | Present, Localized | ||||||
| Romania | Present, Widespread | ||||||
| Serbia and Montenegro | Present, Localized | ||||||
| Spain | Present, Localized | ||||||
| Sweden | Present, Localized | ||||||
| Switzerland | Present, Localized | ||||||
| United Kingdom | Present, Localized | ||||||
North America |
|||||||
| Canada | Present | Present based on regional distribution. | |||||
| -Ontario | Present, Localized | ||||||
| -Quebec | Present, Localized | ||||||
| -Saskatchewan | Present, Localized | ||||||
| Costa Rica | Present, Localized | ||||||
| Mexico | Present, Localized | ||||||
| Panama | Present, Localized | ||||||
| Trinidad and Tobago | Present, Widespread | ||||||
| United States | Present | Present based on regional distribution. | |||||
| -California | Present, Localized | ||||||
| -Colorado | Present, Localized | ||||||
| -Connecticut | Present, Localized | ||||||
| -Georgia | Present, Localized | ||||||
| -Idaho | Present, Localized | ||||||
| -Illinois | Present, Localized | ||||||
| -Indiana | Present, Localized | ||||||
| -Kansas | Present, Localized | ||||||
| -Kentucky | Present, Localized | ||||||
| -Louisiana | Present, Localized | ||||||
| -Maryland | Present, Localized | ||||||
| -Michigan | Present, Localized | ||||||
| -Minnesota | Present | ||||||
| -Missouri | Present, Localized | ||||||
| -Nebraska | Present, Localized | ||||||
| -New Jersey | Present, Localized | ||||||
| -New York | Present, Localized | ||||||
| -North Carolina | Present, Localized | ||||||
| -North Dakota | Present, Localized | ||||||
| -Ohio | Present, Localized | ||||||
| -Oklahoma | Present, Localized | ||||||
| -Pennsylvania | Present, Localized | ||||||
| -South Dakota | Present, Localized | ||||||
| -Texas | Present, Localized | ||||||
| -Utah | Present, Localized | ||||||
| -Vermont | Present, Localized | ||||||
| -Washington | Present, Localized | ||||||
| -Wyoming | Present, Localized | ||||||
Oceania |
|||||||
| Australia | Present | Present based on regional distribution. | |||||
| -Queensland | Present, Localized | ||||||
| -South Australia | Present, Localized | ||||||
| -Victoria | Present, Localized | ||||||
| -Western Australia | Present, Localized | ||||||
| New Zealand | Present, Localized | ||||||
| Papua New Guinea | Present, Localized | ||||||
South America |
|||||||
| Argentina | Present, Widespread | ||||||
| Brazil | Present, Widespread | ||||||
| -Goias | Present, Localized | ||||||
| -Mato Grosso do Sul | Present, Localized | ||||||
| -Minas Gerais | Present, Localized | ||||||
| -Para | Present, Widespread | ||||||
| -Parana | Present, Widespread | ||||||
| -Rio de Janeiro | Present, Widespread | ||||||
| -Sao Paulo | Present, Widespread | ||||||
| Chile | Present, Localized | ||||||
| Colombia | Present, Localized | ||||||
| Venezuela | Present, Localized | ||||||
The pathology of experimentally induced rotaviruses in colostrum-deprived, gnotobiotic calves has been described (Mebus et al., 1973). The changes are unremarkable and consists of dehydration, fluid-filled intestinal tract and distension of the abomasums. The microscopic changes consists of shortening of the length of the villi and replacement of the tall columnar villous epithelial cells by cuboidal and squamous cells. Segments of the small intestine may reveal villous fusion, rounded absorptive cells, villous atrophy and exposure of lamina propria. Crypt hyperplasia occurs in response to the loss of columnar epithelial cells from the villi.
Agrawal et al. (2002) studied the pathogenesis and clinical signs in colostrum-deprived day-old calves. The calves were given 5 ml of bovine rotavirus isolate (106 TCID50/ml) orally which resulted to anorexia, dehydration and profuse bright yellow diarrhoea within 18-24 h after infection and which lasted for 4-5 days. Haematological and biochemical studies showed erythrocytosis, leukocytosis, lymphocytosis and increased packed cell volume, haemoglobin concentration, blood urea nitrogen, hyperproteinaemia, hyperkalaemia and hyponatraemia. Serum alkaline phosphatase, AST and ALT values were also increased in infected calves. Pathological lesions in infected calves sacrificed on day 2, 4 and 6 revealed mucosal congestion and catarrhal exudate in the lumen of small intestine upon microscopic examination. The proximal part of small intestine showed necrosis, desquamation of villous epithelium, shortening of villi with congestion and mononuclear cellular infiltration in sub-mucosa. In the middle and distal part of small intestine, the villus epithelium was necrotic and desquamated. Peyer's patches revealed proliferation of lymphoid cells (Agrawal et al., 2002). Calves were infected orally with four cytopathic strains (81/32F, 81/36F, 81/40F, 82/80F) of bovine rotavirus shown to be pathogenic for conventionally reared newborn calves. All became febrile, were depressed and diarrhoeic. Postmortem examination revealed localized lesions of the small intestines, which are considered to be typical of rotavirus infection (Castrucci et al., 1983).
Hines et al. (1995) reported haemorrhagic enteritis with mild hepatomegaly in a 2-week-old emu chick with a mixed infection of E. coli, adenovirus and rotavirus. Microscopic examination showed necrohaemorrhagic enteritis with intralesional intranuclear basophilic viral inclusion bodies in intestinal epithelial cells; splenic lymphoid necrosis and fibrin exudation; hepatocellular vacuolar change; and multiple clusters of small Gram-negative bacilli in the liver, spleen, yolk sac, and intestine.
Loss of villi, hyperaemia, haemorrhages and neutrophil infiltration of the lamina propria, and oedema and hyperaemia in the submucosa were observed in 12 one-day-old colostrum-deprived piglets orally inoculated with MA86 F23 strain of porcine rotavirus (Ding et al., 1991). Five gnotobiotic piglets inoculated orally with porcine rotavirus developed an enteric lesion (Narita et al., 1982). Electron microscopy of the mucosal epithelium 12 h after inoculation showed that the virus penetrates into the absorptive cells between microvilli, possibly by a pinocytic mechanism. Afterwards, virus particles were most often seen within dilated cisternae of the rough endoplasmic reticulum. These infected cells showed a range of changes, such as disruption of the microvilli, loss of cytoplasmic density and deposition of lipid droplets. Subsequently, most of the epithelial cells were desquamated from the villi. The interaction of the virus and intestinal cells indicates that rotavirus is pathogenic to epithelial cells (Narita et al., 1982).
Extrahepatic biliary atresia was simulated in Balb/c-mice infected with rotavirus. The extension and localization of atresia varied from short to interrupted or long-segment atresia, with or without prestenotic dilatation. The gallbladder was small and atretic, or appeared hydropic due to atresia of the common bile duct (Petersen et al., 1998).
Histopathological changes, including villus shortening and fusion, increased vacuolation of epithelial cells, and mononuclear infiltration of the lamina propria, were observed throughout the small intestine between 12 and 120 h after tissue culture-adapted lapine rotavirus, strain ALA, infection in 1-week-old, 1- to 2-month-old, and 11-month-old rabbits (Ciarlet et al., 1998).
The clinical manifestations of rotavirus gastroenteritis are not distinct enough to permit a specific diagnosis, specimens must be examined in the laboratory. Affected children and young animals and birds may stop feeding and have watery diarrhoea and dehydration. Rotavirus may be present in faeces of both healthy and diarrhoeic animals, and this presents problems in interpreting test results and requires identification of the epidemiological factors which may have precipitated the disease in animals in which the viruses are ubiquitous.
The pathological changes in rotaviral infections are unremarkable and consist of dehydration, fluid-filled intestinal tract and distension of the abomasum. The histopathological changes consist of shortening of the length of the villi and replacement of the tall columnar villous epithelial cells by cuboidal and squamous cells. Segments of the small intestine may reveal villous fusion, rounded absorptive cells, villous atrophy and exposure of lamina propria.
The cause of acute diarrhoea in newborn farm animals cannot usually be identified by using clinical signs. The cause needs to be differentiated from all the common bacterial and viral enteropathogens that cause acute, profuse fluid diarrhoea with progressive dehydration.
In calves, rotavirus diarrhoea needs to be differentiated from enteric collibacillosis, coronavirus diarrhoea, cryptsporidiasis and bovine virus diarrhoea [mucosal disease]. Transmissible gastroenteritis, enteric collibacillosis, coccidiosis, haemorrhagic enterotoxaemia due to Clostridium perfringens type C in piglets resemble rotavirus infection. In lambs, enteric collibacillosis, coliform septicaemia and lamb dysentery needs to be differentiated from rotavirus diarrhoea.
Laboratory diagnosis of rotavirus infections requires identifying the virus in faeces or rectal swab specimens or demonstrating a four-fold or greater increase in antibody to a rotavirus antigen between acute- and convalescent-phase sera. In enteritis in children, the most profuse shedding of virus in faecal matter occurs between days 3 and 5 after onset of disease, and the agent is rarely detected after day 8 (Acha and Szyfres, 2003).
Numerous methods to detect rotavirus in stool and rectal swab specimens have been described. The main features of important diagnostic tests are presented below.
Electron microscopy (EM), with the ability to detect non-Group A rotavirus as well as other viral particles, remains the standard diagnostic technique against which other methods are compared. Immuno-electron microscopy (IEM) is one of the most sensitive EM techniques and use of monoclonal antibodies (Mab) in IEM has proved useful in epidemiological studies of rotavirus infection (Bartlet et al., 1987). It is easier to see the virus if it has been concentrated by ultracentrifugation or clumped by IEM using specific antiserum. The number of viruses may rapidly fall below detectable levels and so faecal specimens (20-30 g) should ideally be collected within 48 h of onset of signs, kept cool and sent to the laboratory. The general view is that at least 106 virus particles per mm of specimen are required for detection by EM. However, it is not practical to examine remnants of food because the number of virus particles will be too low to be detected by currently available methods (Richards GP, 1999).
The isolation of rotavirus from the faeces of calves can be improved by treatment of the faeces with trypsin, which enhances the infectivity of the virus for tissue culture. Treatment of the faeces with chymotrypsin improves the detection rate by isolation (Rhodes et al., 1979). Normally, replication of the virus in tissue culture using conventional methods is limited. Freezing and thawing may sometimes destroy the morphological integrity of variance but not the infectivity of the sample. Samples of intestinal mucosa from several sections of the small and large intestine should be submitted chilled for virus detection and possible isolation. Complete intestines need to be sent from dead birds. The culture of human rotavirus is possible in cell-lines such as MA-104 (Albert and Bishop, 1984) and African green monkey kidney cell culture (Vasileva et al., 1987). However, successful isolation of the pathogen from clinical cases was achieved in only 70% of cases.
Polyacrylamide gel electrophoresis (PAGE) detection of rotavirus in stool is a highly specific, sensitive and inexpensive means of virus detection and can detect both Group A and non-Group A rotaviruses. Using PAGE, epidemiological studies can follow specific rotavirus electropherotypes within a population (Bartlet et al., 1987; Chauhan and Singh, 1993). This test has been found to be more reliable than ELISA for testing samples from newborn children, given that the latter method yields a higher rate of false positives in samples from developing countries (Chen et al., 1999). However, PAGE cannot discriminate between various serotypes (Beards, 1982).
Methods that use anti-rotavirus antibodies include enzyme immunoassay (EIA) or enzyme-linked immunosorbent assay (ELISA), radioimmuno assay (RIA), latex particle agglutination (LA), reverse passive haemagglutination assay (RPHA), and immunofluorescense test (IFT), which includes centrifuging of clinical material onto tissue culture cells followed by immunofluorescence. EIA have proved more sensitive than EM but not more than IEM. Other techniques, which are less sensitive than EIA, include double immuno diffusion (DID), counter immunoelectrophoresis (CIE), co-agglutination reaction, immune adherence haemagglutination and other agglutination techniques (Cukor and Blacklow, 1984; Bartlet et al., 1987). However, antibody-dependent assays have certain limitations for detecting non-Group A rotaviruses. The efficacy of Mab-based EIA in the diagnosis of rotavirus serotypes has been found equal to that of reverse transcriptase polymerase chain reaction (Ushijima et al., 1994).
ELISA has become the method of choice in most laboratories, because it is practical, rapid, and efficient and does not require sophisticated laboratory equipment. Commercial ELISA kits are now available. The ELISA is more sensitive and simple than immunoelectro-osmophoresis complement fixation, IFT on inoculated cell cultures or EM. A counterimmuno-electrophoresis test compares favorably with the ELISA test (Ekern et al., 1981), which uses reagents that are both stable and non-radioactive and is ideal for handling large numbers of specimens. The use of ELISA technology provided an efficient and effective means of evaluating the presence of rotavirus antigen in faecal samples and indicates that this procedure is a very useful tool in epidemiological studies, but that other techniques are required to confirm the presence of the virus (Markowska-Daniel et al., 1996). Culture amplification in colon adenocarcinoma cell line (CaCo-2) combined with enzyme immunoassay (Pathfinder ELISA) was developed as a supplementary tool for rotavirus diagnosis(Cumino et al., 1998). This result was confirmed by PAGE and direct electron microscopy (EM), which increased the rates of rotavirus detection up to 100% after the third, and fifth cell passages, respectively. The combined CaCo-2 cell line amplification-immunoassay method proved to be suitable both to evaluate increase in sensitivity of newly developed rotavirus assays and for rotaviral amplification before antigen assays (Cumino et al., 1998).
Dot-blot or dot-hybridization can be another useful epidemiological tool as it can detect specific rotavirus segments (Flores et al., 1983; Krass’Ko et al., 1987). An AP-labeled oligonucleotide probe has been reported for use in a dot blot format to detect as little as 1 ng of rotavirus RNA. By using individual genes as probes, sub-grouping of rotaviruses isolated from humans is now possible.
IFT and peroxodase-antiperoxidase (PAP) are highly specific for detecting rotaviruses in biopsy materials and cell cultures (Graham and Estes, 1979; Bryden et al., 1987). Virus adsorption and elution (viradel) is used for rotavirus detection in large volumes of water (Goyal and Gerba, 1983).
Serological evidence of rotavirus infection can be detected by various techniques, such as ELISA, IFT, neutralization and complement fixation test (CFT). Agar gel immunodiffusion (AGID) and CIE, CFT, RIA, haemagglutination (HA) and haemagglutination inhibition (HI) tests were compared in their efficiency for detecting bovine rotavirus antigens and antibodies (Mohammed et al., 1978). As a test for antigen using hyperimmune serum, CIE was found to have advantages over AGID by being more rapid as well as approximately four times more sensitive, regardless of whether the antigen was of faecal or tissue culture origin. The CF test was more sensitive than either of the immunodiffusion procedures studied for antigen detection, but was more tedious to perform and of limited use as some faecal samples exhibited anti-complementary activity. For measurement of rotavirus antibody, radio-immunoassay (RIA) was the most sensitive technique and the CIEP least sensitive (Mohammed et al., 1978).
The latex agglutination (LA) test is a simple test that uses particles sensitized with antisera to the virus. Despite being as sensitive as EM, it is less specific, and is therefore best used as a screening test (Acha and Szyfres, 2003). LA is available for the rapid diagnosis of porcine rotavirus infections (Sanekata et al., 1991).
The polymerase chain reaction (PCR) to detect rotavirus in infected faeces has been described. The assay is 100,000 times more sensitive than the standard electrophoretype method that is widely used. It also gives a 5000-fold increase in sensitivity over the previously developed hybridization-based assay and does not require the use of radioisotopes (Xu et al., 1990). The amplified product is a full length c-DNA copy of the gene encoding the major neutralization antigen of the virus whose molecular cloning and sequence analysis will allow rapid collection of detailed information on molecular epidemiology (Xu et al., 1990).
Serotype-specific cDNA molecules produced by using reverse transcription-polymerase chain reaction (RT-PCR) have been used to generate gene probes to detect rotaviruses of porcine origin. RT-PCR targeted to the major outer capsid glycoprotein gene (vp7) has been developed to detect rotavirus in faecal samples from humans with a sensitivity of about 2 ng (equivalent to 108 viral genomes). A PCR typing method has also been reported in which each serotype resulted in a characteristically sized fragment.
A nested reverse transcriptase-polymerase chain reaction (RT-PCR) for detecting Group A rotaviruses was developed by Elschner et al. (2002). It is based on a target region in gene segment 6. Rotavirus strains of human, bovine, porcine, canine, feline, equine and ovine origin were examined. A nested RT-PCR product was formed with all strains and faecal samples tested. The detection limit for virus-containing cell culture supernatant was 3 x 10-2 (50% tissue culture infective dose, TCID50) by RT-PCR and 3 x 10-3 TCID50 by nested amplification. The detection limit of the present PCR procedure was approximately 1.6 x 102 TCID50 per g of faeces, and could be increased by one order of magnitude using nested PCR. The present method for detecting and identifying Group A rotaviruses represents a powerful diagnostic tool and was shown to be applicable to rotaviruses of different origin, including from humans (Elschner et al., 2002).
A method employing guanidine isothiocyanate, hydroxyapatite and cetyltrimethyl ammonium bromide for extraction and purification of rotavirus dsRNA from faecal specimens has not only been found to be very efficient and easy to perform but also to preclude the use of toxic substances such as phenol, chloroform and Freon for this purpose (Santos and Gouvea, 1994). It yields RNA free of enzymatic inhibitors, permitting its detection by RT-PCR assays. In addition, it was demonstrated that during initial clarification of the faecal suspension, the pellet must be washed at least twice to avoid massive losses of virus, viral protein, or viral nucleic acid retained in the solid debris (Santos and Gouvea, 1994). RT-PCR-basedmethod for rotavirus genotyping has proven to be a useful toolfor epidemiologicalinvestigations (Baggi and Peduzzi, 2000).
The immunodiffusion test and EM are superior to the fluorescent antibody technique. Solid-phase radioimmunoassay gives comparable results to EM, but requires radioactive, unstable reagents and expensive counting equipment. PCR showed a much higher sensitivity (93%) than the ELISA test (82.6%) (Taniguchi et al., 1992). The latex agglutination (LA) test has been found to be less sensitive but more specific than ELISA. Agglutination had negative predictive values (94%), compared with agglutination and PAGE, but had the lowest positive predictive value (a measure of accuracy, 70%). Agreement with concentrated polyacrylamide gel electrophoresis (CPAGE) was highest for PAGE (94.8%), followed by agglutination (87%) and ELISA (84.4%) (Hammami et al., 1990). The sensitivity of the latex agglutination (LA) test was found to be four times higher than that of the EM method (Sanekata et al., 1991).
| Sign | Life Stages | Type |
|---|---|---|
| Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed | Cattle and Buffaloes|Calf; Other|Juvenile; Pigs|Piglet; Pigs|Weaner; Poultry|Day-old chick; Poultry|Embryo; Poultry|Young poultry; Sheep and Goats|Lamb | Sign |
| Digestive Signs / Bloody stools, faeces, haematochezia | Cattle and Buffaloes|Calf; Sheep and Goats|Lamb | Sign |
| Digestive Signs / Diarrhoea | Cattle and Buffaloes|All Stages; Other|All Stages; Pigs|All Stages; Poultry|All Stages; Sheep and Goats|All Stages | Sign |
| Digestive Signs / Hepatosplenomegaly, splenomegaly, hepatomegaly | Poultry|Day-old chick | Sign |
| Digestive Signs / Inability to open (trismus) and / or close jaw, mouth | Sheep and Goats|Lamb | Sign |
| Digestive Signs / Steatorrhea, fatty stools, faeces | Pigs|Piglet | Sign |
| Digestive Signs / Unusual or foul odor, stools, faeces | Cattle and Buffaloes|Calf; Other|Juvenile; Sheep and Goats|Lamb | Sign |
| Digestive Signs / Vomiting or regurgitation, emesis | Pigs|Piglet; Pigs|Weaner | Sign |
| General Signs / Dehydration | Cattle and Buffaloes|Calf; Other|Juvenile; Pigs|Piglet; Pigs|Weaner; Poultry|Day-old chick; Poultry|Young poultry; Sheep and Goats|Lamb | Sign |
| General Signs / Discomfort, restlessness in birds | Poultry|All Stages | Sign |
| General Signs / Fever, pyrexia, hyperthermia | Other|Juvenile | Sign |
| General Signs / Inability to stand, downer, prostration | Other|Juvenile | Sign |
| General Signs / Increased mortality in flocks of birds | Poultry|All Stages | Sign |
| General Signs / Lack of growth or weight gain, retarded, stunted growth | Cattle and Buffaloes|Calf; Other|Juvenile; Pigs|Piglet; Pigs|Weaner; Poultry|Day-old chick; Poultry|Young poultry; Sheep and Goats|Lamb | Sign |
| General Signs / Sudden death, found dead | Cattle and Buffaloes|Calf | Sign |
| General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift | Cattle and Buffaloes|All Stages; Other|All Stages; Pigs|All Stages; Poultry|All Stages; Sheep and Goats|All Stages | Sign |
| General Signs / Weight loss | Cattle and Buffaloes|Calf; Other|Juvenile; Pigs|Piglet; Pigs|Weaner; Poultry|Day-old chick; Poultry|Young poultry; Sheep and Goats|Lamb | Sign |
| Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless | Cattle and Buffaloes|Calf; Other|Juvenile; Pigs|Piglet; Pigs|Weaner; Poultry|Day-old chick; Poultry|Embryo; Poultry|Young poultry; Sheep and Goats|Lamb | Sign |
| Pain / Discomfort Signs / Colic, abdominal pain | Other|Juvenile | Sign |
| Reproductive Signs / Agalactia, decreased, absent milk production | Cattle and Buffaloes|Cow | Sign |
| Urinary Signs / Proteinuria, protein in urine | Cattle and Buffaloes|Calf | Sign |
Rotaviruses are transmitted by the faecal-oral route. Other routes of transmission, such as waterborne or airborne (respiratory) routes, have also been suggested. From clinical studies, the incubation period of rotavirus diarrhoeal illness was estimated to be less than 48 h. Large numbers of virus particles are shed in the stool following multiplication in epithelial cells of the small intestine. The rotavirus infects mature brush borer villous epithelial cells in the small intestine, and to lesser extent in the large intestine (Woode and Crouch, 1978). The infected cells are sloughed leading to partial villous atrophy and the atrophic villi are rapidly recovered with relatively undifferentiated crypt cells which mature over a few days and lead to healing of the lesion. The net effect of the morphological and functional changes in the intestine is malabsorption resulting in diarrhoea, dehydration, loss of electrolytes and acidosis. A major factor in the pathogenesis of rotavirus infection in newborn farm animals is the amount of colostrum-derived antibody present in the intestinal lumen at the time of viral challenge (Snodgrass and Wells, 1978).
The pathogenesis of rotaviral infection is similar in calves (Woode and Crouch, 1978), lambs, pigs (Torres-Medina A,Underdahl NR, 1980; McAdaragh et al., 1980) and foals. The lesions occurs within 24 h after infection, villous epithelial cells of the small intestine are infected and become detached. Regeneration occurs within 4-6 days after the onset of the diarrhoea.
Affected calves show mild depression, anorexia and may have a mild fever. The naturally occurring disease usually occurs in calves over 4 days of age and is characterized by a sudden onset of a profuse liquid diarrhoea (De Leeuw et al., 1980). The faeces are pale yellow, mucoid and may contain flecks of blood. The susceptibility of gnotobiotic, colostrum-derived, or sucking calves to four bovine rotavirus isolates was found to be age dependent (Tzipori et al., 1981). Calves older than 7 days remained clinically normal, although they excreted virus in their faeces and subsequently developed antibody against the virus. In general, diarrhoea appeared after a rotavirus incubation period of approximately 3 days and was independent of the order in which the two microbial agents were given, the age of the calf, or the level of circulating rotavirus antibodies (Tzipori et al., 1981).
Affected lambs under 3 weeks of age develop a profuse diarrhoea and the case fatality rate is high. Lambs affected with rotavirus diarrhoea showed frothy salivation, inappetence and diarrhoea, some affected lambs died 4-5 h later and surviving lambs gained weight slowly and were prone to secondary infection (Theil et al., 1996).
Rotaviral diarrhoea may occur in nursed piglets from 1 to 4 weeks of age and in pigs after weaning (Chasey et al., 1986). The disease in nursed piglets resembles milk-scours or 3-week scour. Most of the pigs in the litter are affected with a profuse liquid to soft diarrhoea with varying degrees of dehydration. Recovery usually occurs in a few days unless complicated by enterotoxigenic E. coli or unsatisfactory sanitation, overcrowding and poor management. Experimental rotavirus infection in newborn colostrum-deprived, gnotobiotic piglets results in a profuse liquid diarrhoea in 16-24 h following inoculation. The faeces are yellow and vomiting may occur. Dehydration and death may occur in 2-4 days.
Affected foals appear depressed, fail to suck and become recumbent. Body temperature ranges from 39.5 to 41.0°C and respiration may be rapid and shallow. A profuse, foul-smelling fluid diarrhoea commences 4-12 h after the onset of depression and affected foals become very dehydrated.
Rotavirus infections occur characteristically in young animals. The rotavirus is ubiquitous in the environment of domestic animals and has been isolated in outbreaks of diarrhoea in calves (Woode, 1978), pigs (Bohl, 1979), lambs (Snodgrass et al., 1976) and foals (Conner and Darlington, 1980). The intestinal tract is the site of multiplication of rotavirus and virus is excreted only in the faeces. Because rotaviruses are stable in faeces and relatively resistant to commonly used disinfectants it is extremely difficult to prevent gross contamination of animal housing once infection has been introduced. Diarrhoea associated with rotavirus is observed mainly during winter. The mature animal is considered to be the source of infection for the neonate (Woode, 1978).
There are at least four different groups of rotaviruses definable by serology and electrophoretype (Snodgrass et al., 1984). The most clinically significant strains in the animal and human populations belong to Group A. Cross species transmission of rotavirus has been demonstrated experimentally.
The rotavirus-like virus (rotavirus) was first isolated in the USA in 1969 (Mebus et al., 1969) and was thought to be the cause of outbreaks of diarrheoa in beef calves in Nebraska. While the rotavirus has been most commonly associated with outbreaks of diarrhoea in beef calves raised in groups outdoors, it has also been recovered from dairy herds (De Leeuw, 1980). A unique epidemiological characteristic of rotavirus infection in calves is the short-term nature of the immunity provided by colostrum. The virus is excreted by both calves and adult cattle in large numbers (up to 1010 per g of faeces) and excretion may last for several weeks (McNulty, 1978). There is no evidence that the virus crosses the placental barrier and infects the foetus in utero (Deganais, 1981). The prevalence of subclinical infection may be greater than that indicated by isolation of the virus from faeces. Bovine rotavirus strains of varying virulence can cause subclinical or clinical infections (Bridger and Pocock, 1986).
Healthy calves may also shed the virus and, on examination, show microscopic lesions caused by rotavirus infection (Reynolds et al., 1985). It was shown that adult cattle spread the virus during late pregnancy, especially on the day of calving. Newborn calves then probably became infected by the rotavirus and this is the main infection route for the other newborn calves in the farm (Sahna and Alkan, 2003).
The rotavirus has been isolated from the faeces of lambs with diarrhoea under 3 weeks of age (Snodgrass et al., 1976). The disease appears to be sporadic in lambs and no particular epidemiological characteristics have been described. The experimental disease in lambs is mild and characterized by mild diarrhoea, abdominal discomfort and recovery in a few days (Snodgrass et al., 1977). The mortality in lambs is much higher when both the rotavirus and enteropathogenic E. coli are used (Wray et al., 1981). Affected lambs gain weight slowly and are prone to secondary infection (Theil et al., 1996).
In piglets, rotaviral diarrhoea occurs from 1 to 8 weeks of age, but is commonest in pigs, which are weaned under intensive management conditions at 3 weeks of age (Leece and King, 1978). The disease resembles milk-scours, or 3-week scours of piglets. The morbidity may reach 80% and the case fatality rate ranges from 5 to 20% depending on the level of sanitation. Diarrhoea in unweaned piglets 1-3 weeks of age has been associated with a combined infection of rotavirus and Isopora suis (Nilsson et al., 1984).
Rotavirus is of significance in diarrhoeal syndromes in unweaned piglets, alone or in combination with Eschericia coli or other pathogens (Svensmark et al., 1989a). The findings give evidence to suggest that the type of mild diarrhoea in 3-week-old piglets known as steatorrhoea or ‘white scours’ may be associated with rotavirus infection, possibly in combination with Escherichia coli and other agents. The high prevalence in piglets weaned at 2 weeks, plus the higher morbidity and mortality among such piglets sustain the conclusion that piglets should not be weaned before 3 weeks or age of below a body weight of 6-7 kg (Svensmark et al., 1989b). Environment of commercial pig producers is an important source of rotaviral infection for young piglets (Fu et al. 1989).
Rotaviral diarrhoea occurs in foals from 5 to 35 days of age (Tzipori, 1985). Outbreaks of the disease occur on horse farms with a large number of young foals where the population density is high (Tzipori, 1985). Serological surveys indicated rotavirus antibodies in almost all the mares whose foals were infected with the virus (Tzipori, 1985).
Rotaviruses have been often found in sewage samples (Villena et al., 2003) and wastewater samples (Mehnert and Stewien, 1993). Waste water samples examined during autumn and winter showed a higher rate of presence of rotavirus than those collected in spring and summer. These observations corresponded with the seasonal variation of rotaviral diarrhoea (Mehnert and Stewien, 1993). Market lettuce has been shown to be a mechanical vector for rotavirus (Hemandez et al., 1997). Faecal contamination of food and water is the main route of transmission of enteric viruses. Inadequate treatment of water used to irrigate the crops causes contamination of vegetables (Hemandez et al., 1997). The excreta from infected animals is another potential source of contamination. Animal rotaviruses could also potentially be transmitted by food directly. It has been suggested that rotaviruses may also be airborne (Cook et al., 1996).
Diarrhoea of neonatal calves causes major economic loss directly through mortality and therapy and indirectly from poor growth after clinical disease. It has been estimated that neonatal calf diarrhoea accounts for approximately 75% of the mortality of dairy calves under 3 weeks of age (Radostits et al., 1994). Based on incidence data from two sources, rotaviral infections accounted for 3.2% of infections and 9.1% of loss, ranking 3rd after Escherichia coli and coronaviral infections (House, 1978). In one study, cryptosporidial infections (6.5% loss) were estimated to be similar in economic impact to rotaviral infection. Estimates of economic impact of disease agents in calves (and it is likely to be similar in other species) indicate that rotaviral infections have a relatively minor role in causing economic loss in comparison with E. coli and coronaviral infections (House, 1978). Based on data from two studies, the estimated average annual loss from rotaviral infection was US $3.1 million. The average annual loss of calves from rotavirus and other neonatal diseases between 1970 and 1976, was estimated at US $95,500,000/year (House, 1978). Moreover, the possible long-term effects of neonatal diarrhoea on the health and performance of calves that survive clinical episodes could constitute an even greater loss (Waltner-Toews et al., 1986; Warnick et al., 1995).
In children, rotavirus is a significant cause of morbidity and mortality. In Peru, it has been estimated that 1 in 1.6 children will experience an episode of rotavirus diarrhoea in their first 5 years of life, 1 in 9.4 will seek medical care, 1 in 19.7 will require hospitalization, and 1 in 375 will die of the disease (Ehrenkranz et al., 2001). Per year, this represents approximately 384,000 cases, 64,000 clinic visits, 30,000 hospitalizations, and 1600 deaths. The annual cost of medical care alone for these children is approximately US $2.6 million, and that does not take into account the indirect or societal costs of the illness and the deaths. Rotavirus immunization provides the prospect of decreasing the morbidity and mortality from diarrhoea, but a vaccine regimen would have to be relatively inexpensive, a few dollars or less per child. Future cost-effectiveness analyses should explore the total costs (medical as well as indirect or societal) associated with rotavirus diarrhoea (Ehrenkranz et al., 2001).
Diarrhoea of neonatal calves causes major economic loss directly through mortality and therapy and indirectly from poor growth after clinical disease. It has been estimated that neonatal calf diarrhoea accounts for approximately 75% of the mortality of dairy calves under 3 weeks of age (Radostits et al., 1994). Based on incidence data from two sources, rotaviral infections accounted for 3.2% of infections and 9.1% of loss, ranking 3rd after Escherichia coli and coronaviral infections (House, 1978). In one study, cryptosporidial infections (6.5% loss) were estimated to be similar in economic impact to rotaviral infection. Estimates of economic impact of disease agents in calves (and it is likely to be similar in other species) indicate that rotaviral infections have a relatively minor role in causing economic loss in comparison with E. coli and coronaviral infections (House, 1978). Based on data from two studies, the estimated average annual loss from rotaviral infection was US $3.1 million. The average annual loss of calves from rotavirus and other neonatal diseases between 1970 and 1976, was estimated at US $95,500,000/year (House, 1978). Moreover, the possible long-term effects of neonatal diarrhoea on the health and performance of calves that survive clinical episodes could constitute an even greater loss (Waltner-Toews et al., 1986; Warnick et al., 1995).
In children, rotavirus is a significant cause of morbidity and mortality. In Peru, it has been estimated that 1 in 1.6 children will experience an episode of rotavirus diarrhoea in their first 5 years of life, 1 in 9.4 will seek medical care, 1 in 19.7 will require hospitalization, and 1 in 375 will die of the disease (Ehrenkranz et al., 2001). Per year, this represents approximately 384,000 cases, 64,000 clinic visits, 30,000 hospitalizations, and 1600 deaths. The annual cost of medical care alone for these children is approximately US $2.6 million, and that does not take into account the indirect or societal costs of the illness and the deaths. Rotavirus immunization provides the prospect of decreasing the morbidity and mortality from diarrhoea, but a vaccine regimen would have to be relatively inexpensive, a few dollars or less per child. Future cost-effectiveness analyses should explore the total costs (medical as well as indirect or societal) associated with rotavirus diarrhoea (Ehrenkranz et al., 2001).
There is evidence that zoonotic transmission of rotaviruses, or at least rotavirus genes, can occur. The rapid movement of people of diverse populations throughout the world allows the introduction and spread of rotaviruses generated in developing countries by zoonotic transmission and reassortment.
Studies have shown that genes of animal and human rotaviruses have a lesser degree of homology. Alternatively, identification of unusual rotavirus types from human cases resembling strains from animals supports the view that animal rotaviruses can infect humans.
Rotaviruses cause diarrhoea in newborn animals such as calves, mice, piglets, foals, lambs, rabbits, deer, antelopes, apes, turkeys, chickens, goats, kittens and puppies. Human rotavirus has been shown to induce a diarrhoeal illness in certain animals under experimental conditions. The zoonotic potential of rotaviruses has been reviewed by various workers (Malik et al., 1995; Cook et al., 2004).
Rotaviruses are generally species specific, but cross cross-species transmission has been demonstrated experimentally. It is well demonstrated that animals of one species can be infected by rotaviruses that have been isolated from another species, including humans. Human G1P rotavirus strain Wa has been shown to be pathogenic for experimental pigs (Ward et al. 1996) and is currently used as pathogenic challenge in pigs to assess efficacy of potential human rotavirus vaccines (Ward et al. 1996 and Chang et al. 2001)
Rotavirus isolate PP-1 was highly pathogenic to experimental pigs but replicated poorly in experimental cattle (El-Attar et al., 2001). Sequence analysis showed that this G3 rotavirus had porcine NSP4 and VP4 genes but bovine NSP1 gene.
Epidemiologically, there exists evidence for zoonotic transmission of rotaviruses. Human Group A rotavirus strain possessing genes commonly found in animal rotaviruses have been isolated from infected children in both developed and developing countries. Strains such as G3 (found commonly in species such as cats, dogs monkeys pigs, mice, rabbits and horse), G5 (pigs and horses), G6 and G8 (Cattle), G9 (pigs and lambs) and G10 (cattle) have been isolated from the human population through the world (Desselberger et al., 2001). G and P type combinations, which are found in man, have also been found in animal species. G10P [11] was found in cattle in USA and Canada (Luccheli et al., 1994) and in Indian cows and buffaloes (Gulati et al. 1999). G3P[6] and G4P[6] were found in pigs in Poland and the USA (Winiarczy et al. 2002) and G1P[8] and G5P[8] were found in pigs in Brazil (Santos et al. 1999).
A three-week-old baby in an Israeli household that had a young dog <6 month old age) and was infected with an animal rotavirus G3 strain (Nakagomi et al., 1992). In an outbreak of a Group B rotavirus gastroenteritis in Maryland, USA, in 1985 (Eiden et al. 1985), six out of 16 people (adults and children) with gastroenteritis excreted a virus similar to a Group B rotavirus strain commonly found in rats.
A G8 rotavirus, which had widely circulated in newborn infants in India, causing asymptomatic infection, had VP7 and VP4 gene sequences that were identical to those of a bovine rotavirus strain (Das et al.1993).
During a survey of circulating rotavirus strains carried out in the UK between 1995 and 1998, several uncommon strains were identified (Holmes et al., 1999). Reassortment between common human strains could explain the presence of some of these strains, but not all. Some of the strains with unusual G- and/or P-type, such as G1P[9], G3P[6], G3P[9], G8P[8], G9P[6], and G9P[8] could have been the result of zoonotic transmission or of gene transfer by reassortment. Many of these virus types are found circulating in domestic animals and pets (Desselberger et al., 2001).
Close contact between man and domestic animals may promote exposure to rotaviruses. In areas prone to flooding, or with a monsoon climate, this can increase the chances of contact with animal rotaviruses. All farm workers handling livestock, especially young animals, get contaminated continuously with livestock faeces. Smoking also may be another way in which the viruses can be acquired. Clothing and footwear taken into residential accommodation can also be heavily contaminated.
Aerosolized viruses produced from cleaning practices, such as hosing of pens, could also contaminate hair and skin. Dust and effluent may be potential vehicles of transmission of rotavirus between animals and farm workers, and also to others with access farms. Exposure to animal rotaviruses may occur with farm workers handling sucking calves, piglets and lambs. Non-farm workers can also have direct contact with rotavirus-infected livestock. Several studies have indicated infection of humans by direct contact with household pets. Rotavirus Group A G3 strain, identical to a cat rotavirus strain, has been isolated from a child with a pet cat (Nakagomi and Nakagomi, 1989). Several case studies have indicated infection of humans by direct contact with household pets. Rotavirus infections in dogs are commonly sub-clinical (Saif et al., 1994). Human rotavirus has been shown, experimentally to infect dogs (Tzitori and Makin, 1978). Dogs are also susceptible to infection with porcine (Osterhaus et al., 1980) and bovine rotaviruses (Dagenais et al., 1981). Sequence analysis of VP7 genes from some canine and feline rotavirus strains revealed a high degree of homology between them and may reflect interspecies transmission (Nakagomi et al., 1990).
The excreta from infected cattle, pigs and sheep contain large numbers of infectious rotavirus particles. This is a potential source of contamination in various ways. Viruses in excreta deposited in fields could pass via run-off water into fresh waters such as rivers or lakes. Aerosolized virus could be produced through disturbance of excreta, during cleaning of premises, for example (Cook et al., 2004).
It can be reasonably expected that exposure to animal rotavirus occurs through similar contact, and it would be interesting to examine associated rotavirus incidence. Furthermore, the potential also exists for contamination of watercourses and food crops with animal rotaviruses, via excrement.
There are several enteric viruses, which have been proven transmissible by water (Bosch 1998) and water has been the vehicle in a number of outbreaks of rotavirus gastroenteritis worldwide. A risk of contamination of watercourses with animal rotaviruses exists through animal excreta. Humans could become exposed to animal viruses through drinking untreated water either directly or after failure of a portable water treatment or distribution system. Bathing or recreational water would be another likely source of exposure (Cook et al., 2004).
Animal rotaviruses could potentially be transmitted by foods directly, i.e. through consumption of meat from infected animals, or indirectly, through consumption of food contaminated with organic wastes. Another indirect route of transmission might be handling of food, such as fresh produce, sold on farms by farm workers exposed to infected animals.
The foods which are normally associated with transmission of enteric viral disease are those which are eaten raw, such as soft fruit and salad vegetables, or only lightly cooked, such as shellfish (Svensson, 2000; Cook, 2001). Post cooking contamination of food by improper hygienic practice (e.g. contact with raw pork on preparation surfaces) can be a possibility for transmission. Therefore, foods that have been contaminated by infected handlers after cooking, can also cause outbreaks of viral disease. Group C rotaviruses were the cause of large foodborne outbreak affecting more than 3000 individuals including school children and teachers in seven alimentary schools in Fukui city, Japan (Matsumotto et al., 1989). No particular causative food item was identified through questionnaire responses and the virus could not be detected in food samples. However, the rotavirus isolated from patients was immunologically similar to porcine Group C rotaviruses (Matsumoto et al., 1989).
There is a potential risk of transmission of animal rotaviruses if crops exposed to farmyard waste are eaten raw. It is well known that zoonotic agents can be transmitted via meat contaminated at slaughter. A rating scheme prioritizing the public health significance and potential transmission of zoonotic agents from beef (Petersen et al., 1996) ranked rotavirus 15th in a list of 25 pathogens.
Many viral infections can be spread through air (Sattar and Ijaz, 1987) and it has been suggested that rotaviruses may be airborne (Cook et al., 1996). Spread of rotavirus by airborne droplet has been suggested. Low relative humidity promotes the formation of aerosols. The handling of rotavirus-contaminated material can create aerosols of various sizes. The larger of these settle out rapidly and can contaminate the immediate surroundings, while the smaller (visually <5 m m in diameter) become droplet nuclei, which may remain airborne for a long time (Flewtt 1982; Brandt et al., 1982). Potential routes of airborne exposure to animal rotaviruses exist. Air sucked out either through the side or through the roof of animal housing could result in dissemination of aerolized rotavirus, to which workers and other people living locally could be exposed.
Virus contamination of various objects and surfaces can occur either directly by contact with faeces or indirectly through virus-contaminated aerosols (Sattar and Ijaz, 1987). Ansari et al. (1988) demonstrated potential transfer of rotavirus between surface and hands. It is quite possible that occupational exposure to rotavirus infected animals could be a source of household contamination (Cook et al., 2004).
The primary aim of treatment of rotavirus enteritis is the replacement by the intravenous or oral route of fluids and electrolytes lost by diarrhoea. Withholding of milk for 24-48 h is beneficial. Anti-microbial agents are used both orally and parenterally to treat the possible presence or occurrence of enteric and systemic bacterial infections.
In man, oral rehydration is more readily available and has gained widespread use worldwide as a life-saving treatment. In patients with severe dehydration and shock, intravenous rehydration is indicated for efficient replacement of fluid loss. Use of antiviral drugs like vidarabin, cordycepin and foscarmet for the specific treatment of rotavirus infection have shown promising results in some animal experiments. Of these, foscarmet has been found to be less toxic with high antiviral activity (Rios et al., 1995). Cow colostrum that contains rotavirus antibody, when given prophylactically to infants and young children daily as a single or divided dose orally, was shown to induce significant protection against rotavirus diarrhoea. Oral administration of preparations, which contained rotavirus antibody, has yielded conflicting results with regard to their effectiveness as a method of treatment for children with rotavirus gastroenteritis. Oral bismuth subsalicylate (BSS) as an adjunct to rehydration therapy was reported to shorten the course of rotavirus diarrhoea in a paediatric study.
The problems of rotavirus infection in developed countries contrasts with those in developing countries. In developed countries, mortality is negligible and the aim of rotavirus vaccine is to prevent the infections causing morbidity, whereas in the developing countries, the primary aim is to reduce mortality and morbidity caused by the disease. The rotaviruses are highly contagious and can spread by the faecal-oral route, therefore, careful attention to hand washing, disinfection, and disposal of contaminated material may limit its spread.
Infected young children are the major source of rotavirus infection in the community as they excrete about 1011 virus particles per g of faeces (Flewett, 1985), and the faecal oral-route is the main mode of transmission. Therefore prevention of the disease in children must be based on education and observation of the basic rules of personal hygiene. As with all water-or food-borne diseases, it is important to promote environmental sanitation. In hospitals and nurseries, good hygiene practices are essential. Nosocomial outbreaks of rotavirus infection have been controlled successfully only if the affected wards were closed and staff movement restricted for 10 days (Flewett, 1985). Several studies have shown that breast-fed babies have a lower incidence of disease than bottle fed babies. Rotaviruses have relatively high resistance to chlorine and other common chemical disinfectants. However, 5 mM ethylenediaminetetraacetic acid, ethylene glycol, hydrochloric acid, isopropyl alcohol, glutaraldehyde, hexachloriphene or povidone-iodine can destroy the virus (Acha and Szyfres, 2003).
In animals, rotaviral diarrhoea occurs mainly during the first days of life before there has been time for young animals to become actively immunized. Therefore, investigators have focused their attention on passive protection. Ingestion of maternal colostrum is not always sufficient to prevent disease. To be effective, colostrum has to posses a high antibody titre, as was demonstrated in an experiment that vaccinated cows with a modified virus vaccine with incomplete Freund’s adjuvant (Saif et al., 1983). There are now several vaccines available on the market for cattle and pigs; for cattle, there is an attenuated vaccine that is administered parenterally; for pigs two kinds of vaccines are used, one inactivated and the other live attenuated. The modified live vaccine is given orally to sows three to five weeks before they farrow and once again intramuscularly one week beforehand. The vaccine is also given to sucking pigs 7 to 10 days before weaning. The inactivated vaccine is administered intramuscularly to sows before they farrow and to sucking pigs by the parenteral route (Paul and Lyoo, 1993). These vaccines build up antibody levels in the colostrum and milk of the cow and the sow, and they prolong the excretion of antibodies (Acha and Szyfres, 2003). The main protective antibody is IgA secreted from the intestine. It has been proposed to give colostrum supplements in milk during the period of risk. The management of pregnant animals at the time of parturition must ensure the minimum exposure of newborn animals to infectious agents. Management of the environment of the calf should be emphasized. Stimulation of active immunity by vaccinating the newborn calf with an oral vaccine or stimulation of lactogenic immunity by vaccinating the dam during pregnancy is essential.
Immunity to rotavirusesThe immunological correlates of protection against rotavirus diarrhoea are not well understood (Bishop et al., 1990; Kapikian and Chanock, 1990; Matsui et al., 1989). The role of locally produced intestinal antibody in resistance to illness or infection has not been clearly established, but antibody passively administered via the alimentary tract has been shown to alter susceptibility to disease. In studies of newborn calves, lambs, and mice, the presence of colostrum- or serum-derived rotavirus antibody in the gut lumen at the time of challenge conferred protection against disease whereas circulating antibody did not (Bridger and Woode, 1975; Snodgrass and Wells, 1976).
Antibodies to a variety of rotavirus antigens develop during infection; neutralizing antibodies to both major neutralizing proteins, VP4 and VP7, have been shown to induce homotypic protection in animal studies (Estes, 1990; Offit et al., 1986). However, heterotypic immunity has also been demonstrated in several animal models (Bishop et al.,1986; Wyatt et al., 1975; Zissis et al., 1983). Cell-mediated immunity appears to be involved in protection against rotavirus gastroenteritis in mice (Dharakul et al., 1990; Offit and Dudzik, 1990).
Rotavirus infection of the intestinal enterocytes is thought to be controlled primarily by antibodies. In mice, the appearance of anti-rotaviral IgA in the intestine at 7 days post-infection correlates with the clearance of a primary rotaviral infection (Burns et al., 1995). However, mice lacking in IgA still mount a successful immune response to the pathogen, thought to be mediated by IgG (O'Neal et al., 2000).
CD4+ helper T cells also play a vital role in the successful clearance of rotaviral infection. VanCott et al. (2001) demonstrated that CD4+ T cells are vital to the induction of proper B cell response to rotavirus. Mice lacking CD4+ T cells chronically shed virus in their stool when infected with rotavirus, and produce only 5% of the amount of viral-specific IgA found in healthy mice. In contrast, mice lacking CD8+ T cells responded normally to infection.
It has been shown by Franco and Greenberg that a CD8+ cytotoxic T-lymphocyte (CTL) response is not necessary for clearance of a rotaviral infection (Franco and Greenberg, 1995).
Though CTLs are not necessary, they still play an important role in the development of protective immunity. It was originally observed that severe combined immunodeficient (SCID) mice, which lack both functional B and T cells, develop a chronic rotavirus infection when inoculated with the virus. It was then demonstrated that these mice would clear the infection when CD8+ T cells were transferred from previously vaccinated mice (Dharakul et al., 1990).
It appears that both B and T cells play important roles in the immune response to rotavirus. B cells are involved in the helper T cell-dependent secretion of rotavirus-specific IgA and IgG, while CTLs play a role in the clearance of the virus. The immune system appears to be fully capable of both clearing a rotavirus infection in the absence of either one of these arms of the immune response.
A natural immune state to rotavirus does not exist. Though primary infection by the virus induces production of rotavirus-specific memory B and T cells, these are not normally sufficient to prevent reinfection by the virus. However, they do serve to reduce the severity of secondary infections. It was shown that serum IgA antibody titres correlate with protection against reinfection (Velazquez et al., 2000). It has been shown in mice that in the absence of IgA, IgG is also sufficient to protect mice (O'Neal et al., 2000). One reason these antibody responses do not confer full protection is that they are serotype specific. It has been shown that high amounts of cross reactive secretory IgA, and serotype specific serum IgA and IgG seem to confer the most protection (Anderson and Weber, 2004).
An understanding of the immune response to rotavirus is needed to develop effective prophylaxis. There is evidence that cell-mediated responses may be involved. In experimentally infected calves, most BoCD8+ and BoWC1+ lymphocytes were found in the epithelium, whereas, in control calves, BoCD4+ lymphocytes were predominantly in the lamina propria. The timing and location of these increases in T lymphocyte subsets is indicative of a specific immune response involving BoCD8+ and BoWC1+ T lymphocytes (Parsons et al., 1993).
ImmunizationThe observations made during epidemiology studies throughout the world indicate the need to prevent rotavirus diseases (Snyder and Mershom 1982, Woode, 1978, DeZoysa and Feachem, 1985). Therefore, the WHO working group on diarrhoeal diseases gave considerable priority to the development of rotavirus vaccine to control and prevent rotavirus diarrhoea (Tursi et al., 1989). Significant progress has been made towards the development of a live oral vaccine with the advent of techniques to grow human rotavirus (HRV) in cell culture. The vaccine strains of animal origin are the basis of currently used vaccines for human use as they share the group-specific antigen of HRV, thereby affording protection against HRV or attenuated animal rotavirus (ARV) or reassortants between HRV and ARV. Animals are protected against rotavirus infection by feeding on high-titre anti-rotavirus colostrum (Tsunemitsu et al., 1989). Rotavirus vaccine development has focused on the delivery of live, attenuated rotavirus strains by the oral route.
In humans, the greatest incidence of disease occurs after the age of 5 to 6 months, which allows time for active immunization to take place. Though the first rotavirus vaccine, tetravalent rhesus rotavirus vaccine (RRV-TV) was licensed for human use in the USA in 1998 (Breese et al., 1999), vaccines for human use are still in the stage of development and evaluation (Acha and Szyfres, 2003). Bovine vaccine RIT 4237 was withdrawn from the market, while bovine WC-3 gave good results in trails conducted in USA and is being evaluated in other countries. A vaccine developed with a simian rotavirus strain, rhesus MMU 18006 (or RRV-1), has afforded good protection in controlled trials in Sweden, USA and Venezuela, but it failed in other trials in Finland and USA, probably because it was being tested against a different serotype. Another attempt to develop a vaccine for humans involves a recombinant version of bovine vaccine WC-3 incorporating a human serotype, which is currently undergoing evaluation. There is also a recombinant simian-human vaccine, in which the VP7 antigens from human serotypes 1 and 2 have been incorporated into the simian vaccine rhesus MMU 18006. Trials conducted in Finland and USA showed protection levels of 88% and 67%, respectively, against rotaviral diarrhoea serotype 1 (Acha and Szyfres, 2003). Several live oral rotaviruses have been tested in field trails and have demonstrated an efficacy of 80% or more against severe rotavirus infections with minimal adverse side effects. Ironically, in the past, rotavirus vaccine that demonstrated high efficacy in developed countries appeared to have lowered efficacy in developing countries. This highlights the needs for further evaluation (Breese et al., 1999).
The initial development of rotavirus vaccines for veterinary use was based on Edward Jenner’s approach to smallpox vaccination, which involved the use of an antigenically related, live virus derived from a non-human host. The fact that human and animal rotavirus strains had a common group antigen and that heterologous protection had been demonstrated in animal models supported this approach (Wyatt et al., 1975; Zheng et al., 1988). Gnotobiotic calves that had been immunized with bovine rotavirus strain-Nebraska calf diarrhoea virus (NCDV) in utero did not become ill when challenged at birth with a virulent human rotavirus strain, whereas most control calves developed diarrhoea (Wyatt et al., 1975). A later study showed that gnotobiotic piglets infected with NCDV had a significant reduction in shedding upon subsequent challenge with various human rotavirus strains (Zissis et al., 1983).
The bovine rotavirus vaccine strain RIT4237 was derived from the Lincoln isolate of bovine rotavirus NCDV (Mebus et al., 1971), which bears G6 and P6 serotype specificity; thus, neither of its major neutralization proteins, VP7 or VP4, is shared with the epidemiologically important human rotavirus strains. The vaccine strain RIT4237 was cloned after 147 passages in fetal bovine kidney (FBK) cells, and after 149 passages, the virus was grown in Cercopithecus monkey kidney cells up to passage level 154 (Delem et al., 1984).
Bovine rotavirus vaccine strain WC3 was isolated from the diarrhoeal stool of a newborn calf in Pennsylvania. It was passaged once in African green monkey kidney (AGMK) cells and three times in CV1 cells, plaque purified twice in MA104 cells, and then serially passaged in CV1 cells up to passage level 12 (Clark et al., 1986). Strain WC3 belongs to VP7 serotype 6, as does bovine rotavirus NCDV. The vaccine was administered at a dose of 107 to 107.5 p.f.u. in the efficacy studies. This vaccine was shown to be safe, non-reactogenic, and immunogenic in infants 5 to 11 months of age (Clark et al., 1986).
Rhesus rotavirus (RRV) vaccine strain MMU18006, which was isolated from the diarrhoeal stool of a rhesus monkey, was passaged nine times in primary or secondary monkey kidney cells and then seven times in semicontinuous diploid fetal rhesus lung (DBS-FRhL2) cells (Kapikian et al., 1985). RRV belongs to VP7 serotype 3, one of the epidemiologically important human serotypes, but its VP4 (serotype 5B) is unrelated to that of human strains.
The human-rhesus rotavirus reassortant vaccines were developed with the goal of combining the specificity of the epidemiologically important VP7 serotypes with the attenuation phenotype of RRV (Midthun et al., 1985; 1986). Human-RRV reassortant strains were recovered after co-infection of AGMK cell cultures with the RRV vaccine strain and human rotavirus strain D, DS-1, or ST3. The D (VP7 serotype 1) and DS-1 (VP7 serotype 2) strains were initially detected in stools of children hospitalized with diarrhoea and were then passaged in gnotobiotic calves, whereas ST3 (VP7 serotype 4) was derived from the stool of an asymptomatic newborn infant and was isolated in AGMK cells.
In human-bovine rotavirus reassortant WI79-9, the gene segment encoding theVP7 protein is derived from human rotavirus WI79, a VP7 serotype 1 strain, and the remainder of its genes is derived from bovine rotavirus strain WC3 (Clark et al., 1990).
The initial Jennerian approach involving bovine (RIT4237, WC3) or RRV rotavirus vaccine candidates showed that these vaccines were safe, well tolerated, and immunogenic; however, RRV was more reactogenic than the bovine strains, and in studies in which it was directly compared with RIT4237, it was more immunogenic (Santosham et al., 1991; Vesikari et al., 1986). The rotavirus vaccines of animal origin were similar in that they induced highly variable rates of protection against rotavirus diarrhoea caused by heterotypic strains. In those trials that demonstrated vaccine efficacy, there was generally greater protection against more severe cases of rotavirus diarrhoea; rotavirus infection was not prevented by vaccination. It also appeared that vaccination was more likely to induce heterotypic protection in older infants or those primed by previous infection, perhaps because neutralizing-antibody responses in young infants are predominantly homotypic.
Other approaches to the development of human rotavirus vaccines include cold adaptation of human rotavirus strains or of reassortants between human rotaviruses. A VP7 serotype 1 human rotavirus that was adapted to grow at 26°C was described initially (Matsumo et al., 1987). More recently, cold-adapted reassortant viruses that belong to VP7 serotypes 1 and 2 and reassortants whose VP7 serotype 2 or 3 gene is derived from human rotavirus DS-1 or P, respectively, and whose remaining genes, including the gene encoding VP4 serotype 1A, are derived from human rotavirus Wa have been generated (Hoshimo and Kapikian, 1994; Ostlund et al., 1993).
No vaccine candidates based on recombinant technology have been developed yet for human use. Attempts to express recombinant rotavirus proteins have focused on VP7 and VP4 because of their importance in inducing both passive and active immunity in animal models. VP7 or VP4 genes of simian, bovine, and human rotavirus strains have been expressed by using different bacterial and viral vectors (Andrew et al., 1987; Bellamy and Both, 1990; Estes and Cohen, 1989; Mackow et al., 1989; Mackow et al., 1990; Reeves et al., 1990; Salas-Vidal et al., 1990). Most rotavirus recombinants lack or have limited immunogenicity when inoculated into experimental animals, although baculovirus-expressed VP4 and certain vaccinia- rotavirus VP7 recombinants have induced protection in mice (Andrew et al., 1992; Mackow et al., 1990).
It is clear that the goal of a rotavirus vaccine is not to prevent rotavirus infection or mild illness but, rather, to prevent severe illness in both developed and developing countries. This is a realistic goal, because following a naturally occurring rotavirus infection, reinfections are common. However, the consequences of reinfection are less severe than the initial infection (Kapikian, 1994).
In calves, two approaches have been used to prevent rotavirus-associated diarrhoea in calves, first stimulation of active immunity by vaccinating the newborn calf and second, stimulation of passive lactogenic immunity by vaccinating the pregnant dam (maternal vaccination).
A modified live virus vaccine was first released in 1973 in USA for prevention of rotavirus-associated diarrhoea by oral inoculation of the newborn calves. Later, when oral inoculation of newborn calves did not yield promising results under field conditions, live modified or inactivated vaccines were used to inject the pregnant cows to produce lactogenic immunity in calves. Lactogenic immunity to piglets was also achieved by administration of three doses of an attenuated porcine rotavirus vaccine in pregnant sows at two-week intervals before farrowing (Lecee et al, 1976).
Extremely high rotavirus antibody levels in colostrums and milk were archived by parental injection combined with intramammary infusion of pregnant cows with a modified live strain of NCDV (Saif et al., 1983). This vaccine (containing108 p.f.u./ml of NCDV strain) induced mainly IgG1-type antibodies. Feeding of calves with 1% of pooled colostrums supplement from such cows conferred, on challenge, clinical protection and prevented virus shedding.
Vaccination of pregnant dams with commercial vaccines containing live attenuated rota and coronaviruses and K99 antigen of E. coli in a combination of two or more have been found to afford remarkable protection to newborn animals against enteropathogens (Snodgrass, 1986 and Wieda et al., 1987). Oral administration of chicken egg yolk immunoglobulins (Y Ig) from hens immunized with bovine rotavirus given as a regular supplement to newborn calves has been reported to be a clinically justifiable option for controlling the disease in newborn calves, which for various reasons fail to maintain an effective local gut immunity via passive colostrum transfer (Kuroki et al., 1994).
Acha PN; Szyfres B, 2003. Rotaviral gastroenteritis, In: Zoonoses and communicable diseases common to man and animals, Vol. III (Viruses), Pan American Health Organization (PAHO), New York, 286-297.
Agrawal DK; Singh NP; Chauhan RS, 2002. Clinico-pathomorphological studies on rotavirus infection in neonatal calves. Indian Journal of Veterinary Medicine, 22(2):85-88; 15 ref.
Albert MJ; Bishop RF, 1984. Cultivation of human rotaviruses in cell culture. Journal of Medical Virology, 13: 377-383.
Alfieri AF; Alfieri AA; Barreiros MAB; Leite JPG; Richtzenhain LJ, 2004. G and P genotypes of group A rotavirus strains circulating in calves in Brazil, 1996-1999. Veterinary Microbiology, 99:167-173.
Anathan S; Vaidya V; Subramanian S; Thyagarajan SP, 1988. Rotavirus infection in Tamil Nadu, India. Virus Information Exchange Newsletter, 5:133-134.
Anderson EJ; Weber SG, 2004. Rotavirus infection in adults. Lancet Infectious Diseases, 4(2):91-99.
Andrew ME; Boyle DB; Coupar BEH; Reddy D; Bellamy AR; Both GW, 1992. Vaccinia-rotavirus VP7 recombinants protect mice against rotavirus-induced diarrhoea. Vaccine, 10:185-191.
Andrew ME; Boyle DB; Coupar BEH; Whitfield PL; Both GW; Bellamy AR, 1987. Vaccinia virus recombinants expressing the SA11 rotavirus VP7 glycoprotein gene induce serotype-specific neutralising antibodies. Journal of Virology, 61:1054-1060.
Ansari SA; Sattar SA; Spriangthorpe VS; Wells G A and Tostowaryk W, 1988. Rotavirus survival on human hand and transfer or infectious virus or animate and non-porous inanimate surfaces. Journal of Clinical Microbiology, 26:1513-1518.
Arias CF; Lizano M; Lopez S, 1987. Synthesis in Escherichia coli and immunological characterization of a polypeptide containing the cleavage sites associated with trypsin enhancement of rotavirus SA11 infectivity. Journal of General Virology, 68:633-642.
Arias CF; Romero P; Alvarez V; Lopez S, 1996. Trypsin activation pathway of rotavirus infectivity. Journal of Virology, 70:5832-5839.
Armah GE; Steele AD; Binka FN; Esona MD; Asmah RH; Anto F; Brown D; Green J; Cutts F; Hall A, 2003. Changing patterns of rotavirus genotypes in Ghana: emergence of human rotavirus G9 as a major cause of diarrhea in children. Journal of Clinical Microbiology, 41(6):2317-2322.
Badawy AS; Gerba C; Kelly LM, 1985. Survival of rotavirus SA11 on vegetables. Food Microbiology, 2:199-205.
Baggi F; Peduzzi R, 2000. Genotyping of rotaviruses in environmental water and stool samples in southern Switzerland by nucleotide sequence analysis of 189 base pairs at the 5’ end of the VP7 gene. Journal of Clinical Microbiology, 38(10):3681-3685.
Barman NN; Barman B; Sarma DK; Pensaert MB, 2003. Prevalence of rotavirus and porcine epidemic diarrhoea virus antibodies in pigs of Assam, India. Indian Journal of Animal Sciences, 73:576-578.
Barreiros MAB; Alfieri AF; Medici KC; Leite JPG; Alfieri AA, 2004. G and P genotypes of group A rotavirus from diarrhoeic calves born to cows vaccinated against the NCDV (P[1],G6) rotavirus strain. Journal of Veterinary Medicine (Series B), 51(3):04-109.
Bartlet AV, et al. , 1987. Rotavirus gastroenteritis. Annual Review of Medicine, 38:399-415.
Beards GM, 1982. Polymorphism of genomic RNAs within rotavirus serotypes and subgroups. Archives of Virology, 74:65-70.
Bellamy AR; Both GW, 1990. Molecular biology of rotaviruses. Advances in Virus Research, 38:1-43.
Berg H, 1989. The ice cream industry. Mælkeritidende, 102(18):404-407.
Bhat P; Macaden R; Unnykrishna P; Rao HGV, 1985. Rotavirus and bacterial enteropathogen in acute diarrhoeas of young children in Bangalore. Indian Journal of Medical Research, 82:105-109.
Bishop R; Lund L; Cipriani E; Unicomb L; Barnes G, 1990. Clinical, serological and intestinal immune responses to rotavirus infection of humans. In: de la Maza LM, and Peterson EM, eds. Medical Virology. Vol. 9, New York, USA: Plenum Press, 85-109.
Bohl EH, 1979. Rotaviral diarrhoea in pigs: brief review. Journal of American Veterinary Medical Association, 174(6):613-615.
Bosch A, 1998. Human enteric viruses in the water environment: a minireview. International Microbiology, 1:191-196.
Brandão PE; Cortez A; Ferreira F; Ferreira Neto JS; Gregori F; Heinemann MB; Homem; VSF; Rodriguez CAR; Jerez JA, 2002. Seroprevalence of antibodies against group A rotavirus in cattle from a pioneer frontier in Brazilian amazon. Brazilian Journal of Veterinary Research and Animal Science, 39(1/6):18-20.
Brandt CD; Kim HW; Rodriquez WJ; Arrobio JO; Jeffries BC; Parrott R, 1982. Rotavirus gastroenteritis and weather. Journal of Clinical Microbiology, 16:478-482.
Breese JS; Glass RI; Ivanoff B; Gentsch JR, 1999. Current status and future priorities for rotavirus vaccine development, evaluation and implementation in developing countries: A review. Vaccine, 17:2207-2222.
Bridger JC; Woode GN, 1975. Neonatal calf diarrhea: identification of a reovirus-like (rotavirus) agent in faeces by immunofluorescence and immune electron microscopy. British Veterinary Journal, 131:528-535.
Brown DWG; Mathan MM; Mathew M; Martin R; Beards GM; Mathan VI, 1988. Rotavirus epidemiology in Vellore, South India: group, subgroup, serotype and electrophoretype. Journal of Clinical Microbiology, 26:2410-2414.
Bryden AS; Davies HA; Thouless ME; Flewitt TH, 1987. Diagnosis of rotavirus infection by cell culture. Journal of Medical Microbiology, 10: 121-125.
Burns JW; Krishnaney AA; Vo PT; Rouse RV; Anderson LJ; Greenberg HB, 1995. Analyses of homologous rotavirus infection in the mouse model. Virology, 207(1):143-153.
Chabchoub A; Landolsi F; Cherkaoui M; Ezzaouia M; Ghorbel A, 2000. Rotavirus and diarrhoea in Arabian foal in Tunisia. Revue de Medecine Veterinaire, 151(5):409-414.
Chang KO; Vandl OH; Yuan L; Hodgins DC; Saif LJ, 2001. Antibody secreting cell responses to rotavirus proteins in gnotobiotic pigs inoculated with attenuated or virulent human rotavirus. Journal of Clinical Microbiology, 39:2807-2813.
Chen D; Luongo CL; Nibert ML; Patton JT, 1999. Rotavirus open cores catalyze 5_-capping and methylation of exogenous RNA: evidence that VP3 is a methyltransferase. Virology, 265:120-130.
Chueh LL; Chu R; Li WI; Chang WF, 1982. Isolation and identification of swine rotavirus in Taiwan. Chinese Journal of Microbiology and Immunology, 15(3):212-220.
Ciarlet M; Gilger MA; Barone C; McArthur M; Estes MK; Conner ME, 1998. Rotavirus disease, but not infection and development of intestinal histopathological lesions, is age restricted in rabbits. Virology 251(2): 343-360.
Cicirello HG; Bimal KD; Gupta A; Bhan MK; Gentsch JR; Kumar R; Glass RI, 1994. High prevalence of rotavirus infections among neonates born at hospitals in Delhi, India: predispositions of newborns for infection with unusual rotavirus. Pediatrics and Infectious Diseases Journal, 13:720-724.
Clark HF; Borian FE; Plotkin SA, 1990. Immune protection of infants against rotavirus gastroenteritis by a serotype 1 reassortant of bovine rotavirus WC3. Journal of Infectious Diseases, 161:1099-1104.
Clark HF; Furukawa T; Bell LM; Offit PA; Perrella PA; Plotkin SA, 1986. Immune response of infants and children to low-passage bovine rotavirus (strain WC3). American Journal of Disease in Child, 140:350-355.
Conner ME; Darlington RW, 1980. Rotavirus infection in foals. American Journal of Veterinary Research, 41(10):1699-1703.
Conner ME; Matson DO; Estes MK, 1994. Rotavirus vaccines and vaccination potential. Current topics in Microbiology and Immunology, 185:285-337.
Cook N, 2001. Viruses in food. CPD infection, 2:98-101.
Cook N; Bridger J; Kendall K; Gomara MI; El-Attar L; Gray J, 2004. The zoonotic potential of rotavirus. Journal of Infection, 48:289-302.
Cook SM; Glass RI; LeBaron CW; Ho M-S, 1990. Global seasonality of rotavirus infection. Bulletin of the World Health Organization, 68:171-177.
Cukor G; Blacklow NR, 1984. Human viral gastroenteritis. Microbiological Reviews, 48:157-179.
Dagenais L; Pastoret PP; Massip A; Kaeckenbeeck A; Lansival B, 1981. Failure to isolate rotavirus from bovine meconium. Veterinary Record, 108(1):11-13.
Dagenais L; Schwers A; Pastoret P; Chappuis G, 1981. Propagation of bovine rotavirus by dogs. Veterinary Research, 109:187.
Das M; Dunn SJ; Woode GN; Greenberg HB; Durga Rao C, 1993. Both surface proteins (VP4 and VP7) of an asymptomatic neonatal rotavirus strain (I321) have high levels of sequence identity with the homologous proteins of a serotype 10 bovine rotavirus. Virology (New York), 194(1):374-379; 33 ref.
Desselberger U; Iturriza-Gomara M; Gray JJ, 2001. Rotavirus epidemiology and surveillance. In: Chadwick D, Goode J A, eds. Gastroenteritis viruses. New York, USA: Wiley, 82-100.
Dewey C; Carman S; Pasma T; Josephson G; McEwen B, 2003. Relationship between group A porcine rotavirus and management practices in pig herds in Ontario. Canadian Veterinary Journal, 44(8):649-653.
Dharakul T; Rott L; Greenberg HB, 1990. Recovery from chronic rotavirus infection in mice with severe combined immunodeficiency: virus clearance mediated by adoptive transfer of immune CD8+ T lymphocytes. Journal of Virology, 64(9):4375-4382.
Dickson J; Smith VW; Coackley W; Mckean P; Adams PS, 1979. Rotavirus infection of foals in Western Australia. Australian Veterinary Journal, 55(4):207-208.
Divizia M; Gabrieli R; Donia D; Macaluso A; Bosch A; Guix S; Sainchez G; Villena C; Pint A RM; Palombi L; Buonuomo E; Cenko F; Leno L; Bebeci D; Bino S, 2004. Waterborne gastroenteritis outbreak in Albania. Water Science and Technology, 50(1):57-61.
Doan LT; Okitsu S; Nishio O; Pham DT; Nguyen DH; Ushijima H, 2003. Epidemiological features of rotavirus infection among hospitalized children with gastroenteristis in Ho Chi Minh City, Vietnam. Journal of Medical Virology, 69(4):588-594.
Dutta SR; Khalfan SA; Baig BH; Philipose L; Fulayfil R, 1990. Epidemiology of rotavirus diarrhoea in children under five years in Bahrain. International Journal of Epidemiology, 19(3):722-727.
Ehrenkranz P; Lanata CF; Penny ME; Salazar-Lindo E; Glass RI, 2001. Rotavirus diarrhea disease burden in Peru: the need for a rotavirus vaccine and its potential cost savings. Rev Panam Salud Publica, 10(4):240-248.
Eiden J; Vonderfecht S; Yolken RH, 1985. Evidence that a novel rotavirus-like agent of rats can cause gastroenteritis in man. Lancet, 2:8-11.
Eisa; MI; Mohamed AA, 2004. Role of enteric pathogens in enteritis in lambs, goat kids and children and their zoonotic importance. Veterinary Medical Journal Giza, 52(1):41-59.
Ekern LJ; Schipper IA; McMahon KJ, 1981. Neonatal bovine enteritis: detection of rotavirus by counterimmunoelectrophoresis and enzyme-linked immunosorbent assay. Canadian Journal of Comparative Medicine, 45(2):135-139.
England JJ; Poston RP, 1980. Electron microscopic identification and subsequent isolation of a rotavirus from a dog with fatal neonatal diarrhoea. American Journal Veterinary Research, 41(5):782-783.
Erdogan HM; Ünver A; Günes V; Citil M, 2003. Frequency of rotavirus and coronavirus in neonatal calves in Kars District. Kafkas Üniversitesi Veteriner Fakültesi Dergisi, 9(1):65-68; 27.
Esona MD; Armah GE; Steele AD, 2003. Molecular epidemiology of rotavirus infection in Western Cameroon. Journal of Tropical Pediatrics, 49(3):160-163.
Estes MK, 1990. Rotaviruses and their replication. In: Fields BN, Knipe DM, Chanock RM, Hirsch MS, Melnick JL, Monath TP, Roizman B, eds. Virology. 2nd ed. New York, USA: Raven Press, 1329-1352.
Estes MK, 2001. Rotaviruses and their replication. In: Knipe DM, Howley PM, Griffin DE, Lamb RA, Martin MA, Roizman B, Strais SE eds. Fields virology. 4th ed., Philadelphia, USA: Lipincott William and Wilikins: 1747-1785.
Eugster AK; Scrutchfield WL, 1980. Diagnosis of rotavirus infection in foals. Proceedings, 2nd International Symposium of Veterinary Laboratory Diagnosticians, June 24-26, Lucerne, Switzerland, III:396-399.
Fiore L; Greenberg HB; Mackow ER, 1991. The VP8 fragment of VP4 is the rhesus rotavirus hemagglutinin. Virology, 181:553-563.
Flewett TH, 1982. Clinical features of rotavirus infections. In: Tyrrell DAJ, Kapikian A Z, eds. Virus infection of the gastrointestinal tract. NewYork, USA: Marcel Dekker, 125-145.
Flewett TS, 1985. Rotavirus in the home and hospital nursery. British Medical Journal, 287:568-569.
Flores J; Boeggeman E; Purcell RH; Sereno M; Perez I; White L; Wyatt RG; Chanock RM; Kapikian AZ, 1983. A dot hybridization assay for detection of rotavirus. Lancet, 1:555-559.
Franco MA; Greenberg HB, 1995. Role of B cells and cytotoxic T lymphocytes in clearance of and immunity to rotavirus infection in mice. Journal of Virology, 69(12):7800-7806.
Fujian Provincial Institute of Animal Husbandry and Veterinary Medicine, 1981. Preliminary study in infectious diarrhoea of cattle. Chinese Journal of Veterinary Medicine, 7(4):2-4.
Fukai K; Saito T; Inoue K; Sato M, 2004. Molecular characterization of novel P [14], G8 bovine group A rotavirus, Sun9, isolated in Japan. Virus Research, 105(1):101-106.
Gabbay YB; Homem VSF; Munford V; Alves AS; Mascarenhas JDP; Linhares AC; Rácz ML, 2003. Detection of rotavirus in dogs with diarrhea in Brazil. Brazilian Journal of Microbiology, 34:77-80.
Ghosh SK; Naik TN, 1989. Evidence for a new rotavirus subgroup in India. Epidemiology and Infection, 102:523-530.
Goyal SM; Gerba CP, 1983. Viradel method for detection of rotavirus from sea water. Journal of Virological Methods, 7:279-285.
Graham D; Estes MK, 1979. Comparison of methods of immunocytochemical detection of rotavirus infections. Infection and Immunity, 26:686-689.
Gulati BR; Nakagomi O; Koshimura Y; Nakagomi T; Pandey R, 1999. Relative frequencies of G and P type among rotaviruses from Indian diarrheic cow and buffalo calves. Journal of Clinical Microbiology, 37:2074-2076.
Hernandez F; Monge R; Jimenez C; Taylor L, 1997. Rotavirus and Hepatitis A virus in market lettuce (Lactuca sativa) in Costa Rica. International Journal of Food Microbiology, 37(2-3):221-223.
Holmes JL; Kirkwood CD; Gerna G; Clement JD; Rao MR; Naficy AB; Abu-Elyazeed R; Savarino SJ; Glass RI; Gentsch JR, 1996. Characterization of unusual G8 rotavirus strains isolated from Egyptian children. Achieves of Virology, 144:1381-1396.
Hoshino Y; Jones RW; Kapikian AZ, 2002. Characterization of neutralization specificities of outer capsid spike protein VP4 of selected murine, lapine and human rotavirus strains. Virology, 299:64-71.
Hoshino Y; Kapikian AZ, 1994. Selection of cold-adapted (ca) mutants of human rotaviruses that exhibit a varying degree of growth restriction in vitro. Journal of Virology, 68:7598-7602.
Hoshino Y; Wyatt RG; Greenberg HB; Kalica AR; Flores J; Kapikian AZ, 1983. Serological comparison of canine rotavirus with various simian and human rotaviruses by plaque reduction neutralization and heamagglutination inhibition tests. Infection and Immunity, 41(1):169-173.
House JA, 1978. Economic impact of rotavirus and other neonatal disease agents of animals. Journal of American Veterinary Medical Association, 173:573-576.
Hussain M; Seth P; Dar L; Boor S, 1996. Classification of rotavirus into G and P type with specimens from children with acute diarrhoea in New Delhi, India. Journal of Clinical Microbiology, 34:1592-1594.
Ijaz KM; Alharbi S; Uduman SA; Cheema Y; Sheek-Hussen MM; Alkhair ARA; Shalabi AG; Ijaz SS; Bin-Othman SA; Sattar SA; Liddle LF, 1994. Seasonality and prevalence of rotavirus in Al-Ain, United Arab Emirates. Clinical and Diagnostic Virology, 2(6):323-329.
Institute of Medicine, 1986. The prospects of immunizing against rotavirus. In: New vaccine development: diseases of importance in developing countries, Vol. 2, Washington DC: National Academy Press.
Jayaram H; Estes MK; Prasad BVV, 2004. Emerging themes in rotavirus cell entry, genome organization, transcription and replication. Virus Research, 101:67-81.
Jiang BM; McClure HM; Fankhauser RL; Monroe SS; Glass RI, 2004. Prevalence of rotavirus and norwalk virus antibodies in non-human primates. Journal of Medical Primatology, 33(1):30-33.
Kaljot KT; Shaw RD; Rubin DH; Greenberg HB, 1988. Infectious rotavirus enters cells by direct cell membrane penetration, not by endocytosis. Journal of Virology, 62:1136-1144.
Kammen A-van; Kila V; Van-Kammen A, 1979. A survey for rotavirus antibodies in Papua New Guinea cattle. Australian Veterinary Journal, 55(2):89-90.
Kanitz CL; Milne FJ, 1977. Identification of an equine rotavirus as a cause of neonatal foal diarrhoea. Proceedings, 22nd Annual Convention, American Association of Equine Practioners, 155-165.
Kapikian AZ, 1994. Rhesus rotavirus-based human rotavirus vaccines and observations on selected non-Jennerian approaches to rotaviruses vaccination. In: Kapikian AZ, ed. Viral infections of the gastrointestinal tract. 2nd ed. New York, USA: Marcel Dekker Inc. 443-470.
Kapikian AZ; Chanock RM, 1990. Rotaviruses, In: Fields BN, Knipe DM, Chanock RM, Hirsch MS, Melnick JL, Monath TP, Roizman B, eds. Virology. 2nd ed. New York, USA: Raven Press, 1353-1404.
Kapikian AZ; Midthun K; Hoshino Y; Flores J; Wyatt RG; Glass RI; Askaa J; Nakagomi O; Nakagomi T; Chanock RM; Levine MM; Clements ML; Dolin R; Wright PF; Belshe RB; Anderson EL; Potash L, 1985. Rhesus rotavirus: a candidate vaccine for prevention of human rotavirus disease. In: Lerner R, Brown F, Chanock RM, ed. Vaccines 85. Modern approaches to vaccines: molecular and chemical basis of resistance to viral, bacterial, and parasitic diseases. New York, USA: Cold Spring Harbor, 357-367.
Kelkar SD; Zade JK, 2004. Group B rotaviruses similar to strain CAL-1 have been circulating in Western India since 1993. Epidemiology and Infection, 132(4):745-749.
Kim YH; Lee OS; Choi CU; An SH, 1987. Epidemiological studies on porcine rotavirus infection in piglets. Research Reports of the Rural Development Administration , Livestock & Veterinary, Korea Republic, 29(1):129-133; 18 ref.
Krass’Ko AG, et al. , 1987. Diagnosis of human rotavirus infection by blot hybridization. Voprosy Virusoly, 2:208-212.
Kuroki M; Ohta M; Ikemori Y; Peralta RC; Yokoyama H; Kodama Y, 1994. Passive protection against bovine bovine rotavirus in calves by specific immunoglobins from chickens egg yolk. Archives of Virology, 138:143-148.
Leece JG; King MW, 1978. Role of rotavirus (reo-like) in weanling diarrhea of pigs. Journal of Clinical Microbiology, 8(4):454-458.
Leeuw PW de; Ellens DJ; Straver PJ; van Balken JA; Moerman A; Baanvinger T, 1980. Rotavirus infections in calves in dairy herds. Research in Veterinary Science, 29(2):135-141.
Liprandi F; Gerder M; Bastidas Z; Lopez JA; Pujol FH; Ludert JE; Joelsson DB; Ciarlet M, 2003. A novel type of VP4 carried by a porcine rotavirus strain. Virology, 315:373-380.
Mackow ER; Barnett JW; Chan H; Greenberg HB, 1989. The rhesus rotavirus outer capsid protein VP4 functions as a hemagglutinin and is antigenically conserved when expressed by a baculovirus recombinant. Journal of Virology, 63:1661-1668.
Mackow ER; Vo PT; Broome R; Bass D; Greenberg HB, 1990. Immunization with baculovirus-expressed VP4 protein passively protects against simian and murine rotavirus challenge. Journal of Virology, 64:1698-1707.
Malik SVS; Kumar A; Bhilegaonkar KN, 1995. Rotavirus an emerging enteropathogen of man and animals: an overview. Journal of Communicable Diseases, 27(4):199-207.
Martella V; Ciarlet M; Camarda A; Pratelli A; Tempesta M; Greco G; Cavalli A; Elia G; Decaro N; Terio V; Bozzo G; Camero M; Buonavoglia C, 2003. Molecular characterization of the VP4, VP6, VP7, and NSP4 genes of lapine rotaviruses identified in Italy: emergence of a novel VP4 genotype. Virology, 314:358-370.
Matsui SM; Mackow ER; Greenberg HB, 1989. Molecular determinant of rotavirus neutralization and protection. Advances in Virus Research, 36:181-214.
Matsumoto K; Hatano M; Kobayachi K; Chasegawa A; Yamazaki S; Nakata S; Chiba S; Kimura Y, 1989. An outbreak of acute gastroenteritis associated with acute rotaviral infection in school children. Journal of Infectious Diseases, 160:611-615.
Matsuno S; Murakami S; Tagaki M; Hayashi M; Inouye S; Hasegawa A; Fukai K, 1987. Cold-adaptation of human rotavirus. Virus Research, 7:273-280.
Mawatari T; Taneichi A; Kawagoe T; Hosokawa M; Togashi K; Tsunemitsu H, 2004. Detection of a bovine group C rotavirus from adult cows with diarrhoea and reduced milk production. Journal of Veterinary Medical Science, 66(7):887-890.
McAdaragh JP; Bergeland ME; Meyer RC; Johnshoy MW; Stotz IJ; Benfield DA; Hammer R, 1980. Pathogenesis of rotaviral enteritis in gnotobiotic pigs: a microscopic study. American Journal of Veterinary Research, 41(10):1572-1581.
McNulty MS, 1978. Rotaviruses. Journal of General Virology, 40(1):1-18.
Mebus CA; Stair EL; Rhodes MB; Twiehaus MJ, 1973. Pathology of neonatal calf diarrhoea induced by a coronavirus-like agent. Vet Pathol. 10:45-64.
Mendez E; Arias CF; Lopez S, 1996. Interactions between the two surface proteins of rotavirus may alter the receptor-binding specificity of the virus. Journal of Virology, 70:1218-1222.
Midthun K; Greenberg HB; Hoshino Y; Kapikian AZ; Chanock RM, 1986. Single gene substitution rotavirus reassortants containing the major neutralization protein (VP7) of human rotavirus serotype 4. Journal of Clinical Microbiology, 24:822-826.
Midthun K; Greenberg HB; Hoshino Y; Kapikian AZ; Wyatt RG; Chanock RM, 1985. Reassortant rotaviruses as potential live rotavirus vaccine candidates. Journal of Virology, 53:949-954.
Mohammed KA; Babiuk LA; Saunders JR; Acres SD, 1978. Bovine rotavirus diagnosis: comparison of various antigens and serological tests. Veterinary Microbiology, 3(2):115-127.
Mohammed MEH; Hart CA; Kadden OR, 2003. Viruses and bacteria associated with neonatal camel calf diarrhoea in Eastern Sudan. Emirates Journal of Agricultural Sciences, 15(1):56-62.
Murphy FA; Fauquet CM; Bishop DHL; Ghabrial SA; Jarvis AW; Martelli GP; Mayo MA; Summers MD, 1995. In: Murphy FA, Fauquet CM et al., eds. Virus Taxonomy: Classification and Nomenclature of Viruses, 5th ed, New York, USA: Springer Verlag, 208-239.
Naik TN; Krishnan T, 1996. Rotavirus vaccine: Current status and future prospects. Indian Journal of Medical Research, 104:76-85.
Narita M; Fukusho A; Shimizu Y, 1982. Electron microscopy of the intestine of gnotobiotic piglets infected with porcine rotavirus. Journal of Comparative Pathology, 92(4):589-597.
Nath G; Singh SP; Sanyal SC, 1992. Childhood diarrhoea due to rotavirus in a country. Indian Journal of Medical Research, 95:259-262.
Ness RK; Gion ER; Ekern LJ; Schipper IA, 1981. The diagnosis of rotaviral infection. North Dakota Farm Research, 38(4):16-17.
Nuytten J; Opdenbosch E van; Muylle E; Oyaert W; Vlaminck K; Debouck P, 1983. Isolation of rotavirus from the faeces of a foal with diarrhoea. Vlaams Diergenteeskundig Tijdschriff, 52 (1):1-6.
O’Neal CM; Harriman GR; Conner ME, 2000. Protection of the villus epithelial cells of the small intestine from rotavirus infection does not require immunoglobulin A. Journal of Virology, 74(9):4102-4109.
Offit PA; Clark HF; Blavat G; Greenberg HB, 1986. Reassortant rotaviruses containing structural proteins VP3 and VP7 from different parents induce antibodies protective against each parental serotype. Journal of Virology, 60:491-496.
Offit PA; Dudzik KI, 1990. Rotavirus-specific cytotoxic T lymphocytes passively protect against gastroenteritis in suckling mice. Journal of Virology, 64:6325-6328.
Osterhaus AD; Grost GA; Wirahadiredja RM; Van den Ingh TS, 1980. Canine viral enteritis: prevalence of parvo, corona and rotavirus infections in dogs in Netherlands. Tijdschrift Voor Diergeneeskunde, 105:18-90.
Ostlund EN; Kapikian AZ; Hoshino Y, 1993. Identification of rotavirus genes associated with (268C) cold-adapted (ca) or non-temperature-sensitive (398C) phenotypes. In: 12th Annual Meeting of American Society for Virology, Washington DC, Abstr. 56-59.
Pearson NJ; Fulton RW; Issel CJ; Springer WT, 1982. Prevalence of rotavirus antibody in chickens and horses in Louisiana, USA. Veterinary Record, 110(3):58-59.
Pearson NJ; Fulton RW; Johnson CA; Woods GN; Breitschwerdt EB, 1980. Prevalence of rotaviral antibodies in Louisiana cattle, dogs and cats detected by an indirect fluorescent antibody test. Proceedings, Annual Meeting of the American Association of Veterinary Laboratory Diagnosticians, 23:129-137.
Petersen C; Grasshoff S; Luciano L, 1998. Diverse morphology of biliary atresia in an animal model. Journal of Hepatology, 28(4):603-607.
Petric M; Middleton PJ; Rapley WA; Mehren KG; Grant C, 1981. A survey of zoo mammals for antibody to rotavirus. Canadian Journal of Comparative Medicine, 45(3):327-329.
Polanco G; González M; Manzano L; Cámara J; Puerto M, 2004. Rotavirus in asymptomatic animals: detection and antigenic classification. Archivos de Medicina Veterinaria, 36:65-70.
Prasad BV; Burns JW; Marietta E; Estes MK; Chiu W, 1990. Localization of VP4 neutralization sites in rotavirus by three-dimensional cryo-electron microscopy. Nature, 343:476-479.
Rai A; Kaur R; Khare S; Kumari S; Kumari S, 1986. Prevalence of Rotavirus diarrhoea in Delhi. Virus Information Exchange Newsletter, 3:13.
Rai J, 1989. Study of viral diarrhoea in India (1980-1989). Virus Information Exchange Newsletter, 6:126-127.
Raj P; Jayashree S; Bahn M K; Svensson L; stingzing G; Kumar R; Prasad AK; Khoshoo V; Arora NK, 1987. RNA electrophoretype of rotavirus excreted by asymptomatic neonates. Indian Journal of Virology, 3:1-11.
Ram S; Khurana S; Khurana SB; Sharma S; Vadhera DV; Broor S, 1990. Bioecological factors and rotavirus diarrhoea. Indian Journal of Medical Research, 91:167-170.
Ramchandran M; Das BK; Vij A; Kumar R; Bhamal SS; Kesari N; Rawat H; Bahl L; Thakur S; Woods PA; Glass RI; Bham MK; Gentsch JR, 1996. Unusual diversity of human rotavirus G and P genotype in India. Journal of Clinical Microbiology, 34:436-439.
Rattan A, 1985. Prevalence of rotavirus gasto-enteritis in India. Virus Information Exchange Newsletter, 2:79.
Reeves PR; Johnson MA; Holmes IH; Dyall-Smith ML, 1990. Expression of rotavirus VP7 antigens in fusions with bacterial proteins. Research Microbiology, 141:1019-1025.
Rhodes MB; Stair EL; McCullough RA; McGill LD; Mebus CA, 1979. Comparison of results using electron microscope, immunodiffusion and fluorescent antibody analyses to detect rotavirus in diarrhoeic fecal samples of calves. Canadian Journal of Comparative Medicine, 43(1):84-89.
Richards GP, 1999. Limitation of molecular biological techniques for assessing the virological safety of foods. Journal of Food Protection, 62:691-697.
Rios M; Munoz M; Spencer E, 1995. Antiviral activity of phosphonoformate and rotavirus transcription and replication. Antiviral Research, 27:71-83.
Rodak L; Smíd B; Nevoránkova Z; Smítalová R; Valícek L, 2004. Verification of sensitivity and specificity of group A rotavirus detection in piglets faeces with monoclonal blocking ELISA methods. Journal of Veterinary Medicine (Series B), 51(4):160-165.
Sahna KC; Alkan F, 2003. Role of pregnancy in epidemiology of rotavirus infection in cattle. Salik Bilimleri Dergisi, Firat Üniversitesi (Veteriner), 17(3):203-209.
Saif LJ; Rosen BL; Parwani AV, 1994. Animal rotaviruses In: Kapikian KZ, ed. Viral infection of gastrointestinal tract. New York, USA: Marcel Dekker, 279-367.
Salas-Vidal E; Plebanski M; Castro S; Perales G; Mata E; Lopez S; Arias CF, 1990. Synthesis of the surface glycoprotein of rotavirus SA11 in the aroA strain of Salmonella typhimurium SL 3261. Research Microbiology, 141:883-886.
Santos N; Gouvea V, 1994. Improved method for purification of viral RNA from fecal specimens for rotavirus detection. Journal of Virological Methods, 46(1):11-21.
Santosham M; Letson GW; Wolff M; Reid R; Gahagan S; Adams R; Callahan C; Sack RB; Kapikian AZ, 1991. A field study of the safety and efficacy of two candidate rotavirus vaccines in a Native American population. Journal of Infectious Diseases, 163:483-487.
Sattar SA; Ijaz MK, 1987. Spread of viral infections by aerosols. CRC Critical reviews on Environmental Control, 17:89-131.
Schlafer DH; Scott FW, 1979. Prevalence of neutralizing antibody to the calf rotavirus in New York cattle. Cornell-Veterinarian, 69(3):262-271.
Schwers A; Hoyois P; Chappuis G; Dagenais L; Pastoret PP, 1984. Propagation of bovine rotavirus by cats and dogs. Annales-de-Recherches-Veterinaires, 13(4):303-308.
Singh A; Pandey R, 1990. Molecular biology and vaccinology of mammalian group A rotaviruses. Indian Journal of Microbiology, 30:233-362.
Singh NB, 1988. Prevalence of rotavirus diarrhoea in Manipur, India. Virus Information Exchange Newsletter, 5:66.
Singh NB; Singh LR; Dev LI; Singh Y; Naik TN, 1988. Further studies on rotavirus diarrhoea in Manipur, India. Virus Information Exchange Newsletter, 5: 99.
Singh V; Broor S; Mehta S; Mehta SK, 1990. Molecular epidemiology of human rotavirus infection in Chandigarh. Indian Journal of Medical Research (A), 91:9-14.
Snodgrass DR; Angus KW; Gray EW, 1977. Rotavirus infection in lambs: pathogenesis and pathology. Archives of Virology, 55(4):263-274.
Snodgrass DR; Nakagomi O; Kaga E; Sarasini A; Gerna G, 1992. Human and bovine serotype G8 rotaviruses may be derived by reassortment. Archives of Virology, 125(1-4):121-128.
Snodgrass DR; Smith W; Gray EW; Herring JA, 1976. A rotavirus in lambs with diarrhoea. Research in Veterinary Science, 20(1):113-114.
Snodgrass DR; Wells PW, 1976. Rotavirus infection in lambs: studies on passive protection. Achieves Virology, 52:201-205.
Snodgrass DR; Wells PW, 1978. Passive immunity in rotaviral infections. Journal of American Veterinary Medical Association, 173(5):565-568.
Snyder JD; Mershon MH, 1982. The magnitude of the global problem of acute diarrhoeal disease: a review of active surveillance data. Bulletin World Health Organization, 60:605-613.
Strickland KL; Lenihan P; O’Connor MG; Condon JC, 1982. Diarrhoea in foals associated with rotavirus (correspondence). Veterinary Record, 111(18):421.
Svensson L, 2000. Diagnosis of foodborne viral infection in patients. International Journal of Food Microbiology, 59:117-126.
Talukder MRI; Huque AKMF; Ahmed MU, 2003. Rotavirus diarrhoea in human and goat kids. Bangladesh Veterinarian, 20:7-12.
Tamehiro CY; Alfieri AF; Medici KC; Alfieri AA, 2003. Segmented double-stranded genomic RNA viruses in fecal samples from broiler chicken. Brazilian Journal of Microbiology, 34(4):349-353.
Torres-Medina A; Underdahl NR, 1980. Scanning electron microscopy of intestine of gnotobiotic piglets infected with porcine rotavirus. Canadian Journal of Comparative Medicine, 44(4):403-411.
Tsai CH; Chiu HH; Abe T, 2000. Epidemiologic features of rotavirus infection in Taiwan: a review. Pediatrics International, 42(4):411-414.
Tsunemitsu H; Imagawa H; Togo M; Shouji T; Kawashima K; Horino R; Imai K; Nishimori T; Takagi M; Higuchi T, 2001. Predominance of G3B and G14 equine group A rotaviruses of a single VP4 serotype in Japan. Archives of Virology, 146(10):1949-1962.
Tsunemitsu H; Shimizu M; Hirai T; Yonemichi H; Kudo T; Mori K; Onoe S, 1989. Protection against rotaviruses in newborne calves by continuous feeding of immune colostrum. Nippon Juigaku Zasshi, 51 (2): 300-308.
Tursi JM; Albert MJ; Bishop RF, 1989. Production and characterization of neutralizing monoclonocal antibody to a human rotavirus strain with a supershort RNA pattern. Journal of Clinical Microbiology, 25:2426-2427.
Tzipori S; Makin T, 1978. Propagation of human rotavirus in young dogs. Veterinary Microbiology, 3:55-63.
Tzipori SR; Makin TJ; Smith ML; Krautil FL, 1981. Clinical manifestations of diarrhea in calves infected with rotavirus and enterotoxigenic Escherichia coli. Journal of Clinical Microbiology, 13(6):1011-1016.
Ushijima H; Mukoyama A; Hasegawa A; Nishimura S; Konishi K; Bosu K, 1994. Serotyping of human rotaviruses in the Tokyo area (1990-1993) by enzyme immuno assay with monoclonal antibodies and by reverse transcription and polymerase chain reaction amplification. Journal of Medical Virology, 44:164-165.
Valenzuela S; Pizarro J; Sandino AM; Vasquez M; Fernandez J; Hernandez O; Patton J; Spencer E, 1991. Photoaffinity labeling of rotavirus VP1 with 8-azido-ATP: identification of the viral RNA polymerase. Journal of Virology, 65:3964-3967.
VanCott JL; McNeal MM; Flint J; Bailey SA; Choi AH; Ward RL, 2001. Role for T cell-independent B cell activity in the resolution of primary rotavirus infection in mice. European Journal of Immunology, 2001, 31(11):3380-3387.
Varghese V; Das S; Singh NB; Kojima K; Bhattacharya SK; Krishnan T; Kobayashi N; Naik TN, 2004. Molecular characterization of a human rotavirus reveals porcine characteristics in most of the genes including VP6 and NSP4. Archives of Virology, 149(1):155-172.
Vasileva VI et al, 1987. Isolation of human rotavirus in a green monkey cell culture. Voprosy Virosily, 3: 360-362.
Velazquez FR; Matson DO; Guerrero ML; Shults J; Calva JJ; Morrow AL; Glass RI; Pickering LK; Ruiz-Palacios GM, 2000. Serum antibody as a marker of protection against natural rotavirus infection and disease. Journal of Infectious Diseases, 182(6):1602-1609.
Vesikari T; Kapikian AZ; Delem A; Zissis G, 1986. A comparative trial of rhesus monkey (RRV-1) and bovine (RIT 4237) oral rotavirus vaccines in young children. Journal of Infectious Diseases, 153:832-839.
Villena C; El-Senousy WM; Abad FX; Pintó RM; Bosch A, 2003. Group A Rotavirus in Sewage Samples from Barcelona and Cairo: Emergence of Unusual Genotypes. Applied and Environmental Microbiology, 69(7):3919-3923.
Wani SA; Bhat MA; Nawchoo R; Munshi ZH; Bach AS, 2004. Evidence of rotavirus associated with neonatal lamb diarrhoea in India. Tropical Animal Health and Production, 36:27-32.
Wani SA; Bhat MA; Samanta I; Ishaq SM; Ashrafi MA; Buchh AS, 2004. Epidemiology of diarrhoea caused by rotavirus and Escherichia coli in lambs in Kashmir valley, India. Small Ruminant Research, 52:145-153.
Whittier CA; Horne W; Slenning B; Loomis M; Stoskopf MK, 2004. Comparison of storage methods for reverse-transcriptase PCR amplification of rotavirus RNA from gorilla (Gorilla g. gorilla) fecal samples. Journal of Virological Methods, 116(1):11-17.
Woode GN, 1978. Epizootiology of bovine rotavirus infection. Veterinary Record, 103(3):44-46.
Woode GN; Crouch CF, 1978. Naturally occurring and experimentally induced rotaviral infections of domestic and laboratory animals. Journal of American Veterinary Medical Association, 173:522-526.
Wray C; Dawson M; Afshar A; Lucas M, 1981. Experimental Escherichia coli and rotavirus infection in lambs. Research in Veterinary Science, 30(3):379-381.
Wyatt RG; Mebus CA; Yolken RH; Kalica AR; James HD Jr; Kapikian AZ; Chanock RM, 1975. Rotaviral immunity in gnotobiotic calves: heterologous resistance to human virus induced by bovine virus. Science, 203:548-550.
Yachha SK; Singh V; Kanwar SS; Mehta S, 1994. Epidemiology, subgroups and serotypes of rotavirus diarrhoea in North Indian communities. Indian Pediatrics, 31:27-33.
Zheng BJ; Han SX; Yan YK; Liang XR; Ma GZ; Yang Y; Ng MH, 1988. Development of neutralizing antibodies and group A common antibodies against natural infections with human rotavirus. Journal of Clinical Microbiology, 26:1506-1512.
Zoysa I de; Feachem RG, 1985. Interventions for the control of diarrheal diseases among young children: rotavirus and cholera immunization. Bulletin of World Health Organization, 63:569-583.
Anathan S, Vaidya V, Subramanian S, Thyagarajan SP, 1988. Rotavirus infection in Tamil Nadu, India. In: Virus Information Exchange Newsletter, 5 133-134.
Bhat P, Macaden R, Unnykrishna P, Rao HGV, 1985. Rotavirus and bacterial enteropathogen in acute diarrhoeas of young children in Bangalore. In: Indian Journal of Medical Research, 82 105-109.
CABI, Undated. Compendium record. Wallingford, UK: CABI
CABI, Undated a. CABI Compendium: Status inferred from regional distribution. Wallingford, UK: CABI
CABI, Undated b. CABI Compendium: Status as determined by CABI editor. Wallingford, UK: CABI
Cicirello HG, Bimal KD, Gupta A, Bhan MK, Gentsch JR, Kumar R, Glass RI, 1994. High prevalence of rotavirus infections among neonates born at hospitals in Delhi, India: predispositions of newborns for infection with unusual rotavirus. In: Pediatrics and Infectious Diseases Journal, 13 720-724.
Dickson J, Smith VW, Coackley W, Mckean P, Adams PS, 1979. Rotavirus infection of foals in Western Australia. In: Australian Veterinary Journal, 55 (4) 207-208.
Hussain M, Seth P, Dar L, Boor S, 1996. Classification of rotavirus into G and P type with specimens from children with acute diarrhoea in New Delhi, India. In: Journal of Clinical Microbiology, 34 1592-1594.
Nath G, Singh SP, Sanyal SC, 1992. Childhood diarrhoea due to rotavirus in a country. In: Indian Journal of Medical Research, 95 259-262.
Pearson NJ, Fulton RW, Johnson CA, Woods GN, Breitschwerdt EB, 1980. Prevalence of rotaviral antibodies in Louisiana cattle, dogs and cats detected by an indirect fluorescent antibody test. [Proceedings, Annual Meeting of the American Association of Veterinary Laboratory Diagnosticians], 23 129-137.
Rai A, Kaur R, Khare S, Kumari S, Kumari S, 1986. Prevalence of Rotavirus diarrhoea in Delhi. In: Virus Information Exchange Newsletter, 3 13.
Rai J, 1989. Study of viral diarrhoea in India (1980-1989). In: Virus Information Exchange Newsletter, 6 126-127.
Raj P, Jayashree S, Bahn MK, Svensson L, Stingzing G, Kumar R, Prasad AK, Khoshoo V, Arora NK, 1987. RNA electrophoretype of rotavirus excreted by asymptomatic neonates. In: Indian Journal of Virology, 3 1-11.
Ram S, Khurana S, Khurana SB, Sharma S, Vadhera DV, Broor S, 1990. Bioecological factors and rotavirus diarrhoea. In: Indian Journal of Medical Research, 91 167-170.
Ramchandran M, Das BK, Vij A, Kumar R, Bhamal SS, Kesari N, Rawat H, Bahl L, Thakur S, Woods PA, Glass RI, Bham MK, Gentsch JR, 1996. Unusual diversity of human rotavirus G and P genotype in India. In: Journal of Clinical Microbiology, 34 436-439.
Rattan A, 1985. Prevalence of rotavirus gasto-enteritis in India. In: Virus Information Exchange Newsletter, 2 79.
Schwers A, Hoyois P, Chappuis G, Dagenais L, Pastoret PP, 1984. Propagation of bovine rotavirus by cats and dogs. In: Annales-de-Recherches-Veterinaires, 13 (4) 303-308.
Singh NB, 1988. Prevalence of rotavirus diarrhoea in Manipur, India. In: Virus Information Exchange Newsletter, 5 66.
Strickland KL, Lenihan P, O'Connor MG, Condon JC, 1982. Diarrhoea in foals associated with rotavirus (correspondence). In: Veterinary Record, 111 (18) 421.
Tsai CH, Chiu HH, Abe T, 2000. Epidemiologic features of rotavirus infection in Taiwan: a review. In: Pediatrics International, 42 (4) 411-414.
| Website | URL | Comment |
|---|---|---|
| Capetown University - rotavirus structure | http://web.uct.ac.za/depts/mmi/stannard/rota.html | Pictures and brief text describing rotavirus structure. |
| Center for Disease Control - rotavirus fact sheet | http://www.cdc.gov/ncidod/dvrd/revb/gastro/rotavirus.htm | Information about rotavirus provided by the US Center for Disease Control. |
| Rotavirus factsheet on www.nlv.com | http://www.nlv.ch/Rotavirus/Rotafactsheet.htm | |
| Rotavirus Vaccine Program - rotavirus facts | http://www.rotavirusvaccine.org/rotavirus-facts.htm | Information on rotavirus provided by Rotavirus Vaccine Program. |
| Virology On-line - rotavirus | http://www.virology-online.com/viruses/Diarrhoea2.htm | Information about rotavirus and other viruses that cause diarrhoea in humans. Site is run by Dr Derek Wong, Hong Kong. |
| WHO - Initiative for Vaccine Research (Diarrhoeal diseases) | http://www.who.int/vaccine_research/documents/new_vaccines/en/index1.html | Site summarizing the WHO's Initiative for Vaccine Research including some basic information on the diseases targeted. This page gives information on diarrhoeal diseases. |
| Wrongdiagnosis.com - rotavirus | http://www.wrongdiagnosis.com/r/rotavirus/intro.htm | Information about rotavirus infections in children. |
One or more of the features that are needed to show you the maps functionality are not available in the web browser that you are using.
Please consider upgrading your browser to the latest version or installing a new browser.
More information about modern web browsers can be found at http://browsehappy.com/
