Antibody (Ig) ELISAs are used to detect historical BVDV infection; these tests have been validated in serum, milk and bulk milk samples. Ig ELISAs do not diagnose active infection but detect the presence of antibodies produced by the animal in response to viral infection. Vaccination also induces an antibody response, which can result in false positive results, therefore it is important to know the vaccination status of the herd or individual when interpreting results. A standard test to assess whether virus has been circulating recently is to perform an Ig ELISA on blood from 5–10 young stock that have not been vaccinated, aged between 9 and 18 months. A positive result indicates exposure to BVDV, but also that any positive animals are very unlikely to be PI animals themselves. A positive result in a pregnant female indicates that she has previously been either vaccinated or infected with BVDV and could possibly be carrying a PI fetus, so antigen testing of the newborn is vital to rule this out. A negative antibody result, at the discretion of the responsible veterinarian, may require further confirmation that the animal is not in fact a PI.

At a herd level, a positive Ig result suggests that BVD virus has been circulating or the herd is vaccinated. Negative results suggest that a PI is unlikely however this naïve herd is in danger of severe consequences should an infected animal be introduced. Antibodies from wild infection or vaccination persist for several years therefore Ig ELISA testing is more valuable when used as a surveillance tool in seronegative herds.

Antigen ELISA and rtPCR are currently the most frequently performed tests to detect virus or viral antigen. Individual testing of ear tissue tag samples or serum samples is performed. It is vital that repeat testing is performed on positive samples to distinguish between acute, transiently infected cattle and PIs. A second positive result, acquired at least three weeks after the primary result, indicates a PI animal. rtPCR can also be used on bulk tank milk (BTM) samples to detect any PI cows contributing to the tank. It is reported that the maximum number of contributing cows from which a PI can be detected is 300.

As in humans, the sensitivity of testing methods for rodents contributes to the accuracy of diagnosis. LCMV is typically identified through serology. However, in an endemically infected colony, more practical methods include MAP (mouse antibody production) and PCR testing. Another means of diagnosis is introducing a known naïve adult mouse to the suspect rodent colony. The introduced mouse will seroconvert, allowing use of immunofluorescence antibody (IFA), MFIA or ELISA to detect antibodies.

Viral disease is usually detected by clinical presentation, for instance severe muscle and joint pains preceding fever, or skin rash and swollen lymph glands.

Laboratory investigation is not directly effective in detecting viral infections, because they do not themselves increase the white blood cell count. Laboratory investigation may be useful in diagnosing associated bacterial infections, however.

Viral infections are commonly of limited duration, so treatment usually consists in reducing the symptoms; antipyretic and analgesic drugs are commonly prescribed.

Immunosuppressive therapy has been effective in halting the disease for laboratory animals.

The MAYV infection is characterized by fever, headache, myalgia, rash, prominent pain in the large joints, and association with rheumatic disease, but these signs and symptoms are unspecific to distinguish from other Arbovirus. The MAYV infection can be confirmed by laboratory testing such us virus isolation, RT-PCR and serology. The virus isolation in cell culture is effective during viremia. RT-PCR helps to identify virus. Serology tests detect antibodies like IgM and the most common assay is IgM-capture enzyme-linked immunosorbant assays (ELISA). This test usually requires a consecutive retest to confirm increasing titers. While the IgG detection is applied for epidemiology studies.

Previous methods of diagnosis included HI, complement fixation, neutralization tests, and injecting the serum of infected individuals into mice. However, new research has introduced more efficient methods to diagnose KFDV. These methods include: nested RT-PCR, TaqMan-based real-time RT-PCR, and immunoglobin M antibodies detection by ELISA. The two methods involving PCR are able to function by attaching a primer to the NS-5 gene which is highly conserved among the genus to which KFDV belongs. The last method allows for the detections of anti-KFDV antibodies in patients.

MVD is clinically indistinguishable from Ebola virus disease (EVD), and it can also easily be confused with many other diseases prevalent in Equatorial Africa, such as other viral hemorrhagic fevers, falciparum malaria, typhoid fever, shigellosis, rickettsial diseases such as typhus, cholera, gram-negative septicemia, borreliosis such as relapsing fever or EHEC enteritis. Other infectious diseases that ought to be included in the differential diagnosis include leptospirosis, scrub typhus, plague, Q fever, candidiasis, histoplasmosis, trypanosomiasis, visceral leishmaniasis, hemorrhagic smallpox, measles, and fulminant viral hepatitis. Non-infectious diseases that can be confused with MVD are acute promyelocytic leukemia, hemolytic uremic syndrome, snake envenomation, clotting factor deficiencies/platelet disorders, thrombotic thrombocytopenic purpura, hereditary hemorrhagic telangiectasia, Kawasaki disease, and even warfarin intoxication. The most important indicator that may lead to the suspicion of MVD at clinical examination is the medical history of the patient, in particular the travel and occupational history (which countries and caves were visited?) and the patient's exposure to wildlife (exposure to bats or bat excrements?). MVD can be confirmed by isolation of marburgviruses from or by detection of marburgvirus antigen or genomic or subgenomic RNAs in patient blood or serum samples during the acute phase of MVD. Marburgvirus isolation is usually performed by inoculation of grivet kidney epithelial Vero E6 or MA-104 cell cultures or by inoculation of human adrenal carcinoma SW-13 cells, all of which react to infection with characteristic cytopathic effects. Filovirions can easily be visualized and identified in cell culture by electron microscopy due to their unique filamentous shapes, but electron microscopy cannot differentiate the various filoviruses alone despite some overall length differences. Immunofluorescence assays are used to confirm marburgvirus presence in cell cultures. During an outbreak, virus isolation and electron microscopy are most often not feasible options. The most common diagnostic methods are therefore RT-PCR in conjunction with antigen-capture ELISA, which can be performed in field or mobile hospitals and laboratories. Indirect immunofluorescence assays (IFAs) are not used for diagnosis of MVD in the field anymore.

Although infection of avian reovirus is spread worldwide, it is rarely the sole cause of a disease. For chickens, the most common manifestation of the disease is joint/limb lameness. Confirming infection of avian reovirus can be detected through an ELISA test by using and observing the expression of σC and σB proteins. However, isolating and identifying reoviruses from tissue samples is very time consuming. Isolation is most successfully attained through inoculation of material into chick embryo cultures or fertile chicken eggs. Inoculation of embryonic eggs through the yolk sac has shown that the virus usually kills the embryos within 5 or 6 days post inoculation. Analyzing the samples, the embryos appeared hemorrhagic and necrotic lesions on the liver were present. (Jones, Onunkwo, 1978). There have also been approaches to identify avian reoviruses molecularly by observing infected tissues with dot-blot hybridization, PCR, and a combination of PCR and RFLP. This combination allows for the reovirus strain to be typed.

Marburgviruses are World Health Organization Risk Group 4 Pathogens, requiring Biosafety Level 4-equivalent containment, laboratory researchers have to be properly trained in BSL-4 practices and wear proper personal protective equipment.

Diagnosis can be made in several ways, encompassing a range of multi-faceted techniques:

- Isolation and detection of the virus in cell culture.

- Detection of viral antigens directly within bodily respiratory tract secretions using immunofluorescence, enzyme immunoassays or fluroimmunoassays.

- Polymerase chain reaction (PCR).

- Analysis of specific IgG antibodies showing a subsequent rise in titre following infection (using paired serum specimens).

Because of the similarity in terms of the antigenic profile between the viruses, hemagglutination assay (HA) or hemadsorption inhibition (HAdI) processes are often used. Both complement fixation, neutralisation and enzyme linked immunosorbent assays – ELISA, can also be used to aid in the process of distinguishing between viral serotypes.

Diagnosis of BMCF depends on a combination of history and symptoms, histopathology and detection in the blood or tissues of viral antibodies by ELISA or of viral DNA by PCR. The characteristic histologic lesions of MCF are lymphocytic arteritis with necrosis of the blood vessel wall and the presence of large T lymphocytes mixed with other cells. The similarity of MCF clinical signs to other enteric diseases, for example blue tongue, mucosal disease and foot and mouth make laboratory diagnosis of MCF important. The world organisation for animal health recognises histopathology as the definitive diagnostic test, but laboratories have adopted other approaches with recent developments in molecular virology. No vaccine has as yet been developed.

Japanese encephalitis is diagnosed by commercially available tests detecting JE virus-specific IgM antibodies in serum and /or cerebrospinal fluid, for example by IgM capture ELISA.

JE virus IgM antibodies are usually detectable 3 to 8 days after onset of illness and persist for 30 to 90 days, but longer persistence has been documented. Therefore, positive IgM antibodies occasionally may reflect a past infection or vaccination. Serum collected within 10 days of illness onset may not have detectable IgM, and the test should be repeated on a convalescent sample. For patients with JE virus IgM antibodies, confirmatory neutralizing antibody testing should be performed.

Confirmatory testing in the US is only available at CDC and a few specialized reference laboratories. In fatal cases, nucleic acid amplification, and virus culture of autopsy tissues can be useful. Viral antigen can be shown in tissues by indirect fluorescent antibody staining.

Chikungunya is diagnosed on the basis of clinical, epidemiological, and laboratory criteria. Clinically, acute onset of high fever and severe joint pain would lead to suspicion of chikungunya. Epidemiological criteria consist of whether the individual has traveled to or spent time in an area in which chikungunya is present within the last twelve days (i.e.) the potential incubation period). Laboratory criteria include a decreased lymphocyte count consistent with viremia. However a definitive laboratory diagnosis can be accomplished through viral isolation, RT-PCR, or serological diagnosis.

The differential diagnosis may include infection with other mosquito-borne viruses, such as dengue or malaria, and infection with influenza. Chronic recurrent polyarthralgia occurs in at least 20% of chikungunya patients one year after infection, whereas such symptoms are uncommon in dengue.

Virus isolation provides the most definitive diagnosis, but takes one to two weeks for completion and must be carried out in biosafety level III laboratories. The technique involves exposing specific cell lines to samples from whole blood and identifying chikungunya virus-specific responses. RT-PCR using nested primer pairs is used to amplify several chikungunya-specific genes from whole blood, generating thousands to millions of copies of the genes in order to identify them. RT-PCR can also be used to quantify the viral load in the blood. Using RT-PCR, diagnostic results can be available in one to two days. Serological diagnosis requires a larger amount of blood than the other methods, and uses an ELISA assay to measure chikungunya-specific IgM levels in the blood serum. One advantage offered by serological diagnosis is that serum IgM is detectable from 5 days to months after the onset of symptoms, but drawbacks are that results may require two to three days, and false positives can occur with infection due to other related viruses, such as o'nyong'nyong virus and Semliki Forest virus.

Presently, there is no specific way to test for chronic signs and symptoms associated with Chikungunya fever although nonspecific laboratory findings such as C reactive protein and elevated cytokines can correlate with disease activity.

The above signs, especially fever, respiratory signs, neurological signs, and thickened footpads occurring in unvaccinated dogs strongly indicate canine distemper. However, several febrile diseases match many of the signs of the disease and only recently has distinguishing between canine hepatitis, herpes virus, parainfluenza and leptospirosis been possible. Thus, finding the virus by various methods in the dog's conjunctival cells or foot pads gives a definitive diagnosis. In older dogs that develop distemper encephalomyelitis, diagnosis may be more difficult, since many of these dogs have an adequate vaccination history.

An additional test to confirm distemper is a brush border slide of the bladder transitional epithelium of the inside lining from the bladder, stained with Dif-Quick. These infected cells have inclusions which stain a carmine red color, found in the paranuclear cytoplasm readability. About 90% of the bladder cells will be positive for inclusions in the early stages of distemper.

Shingles can be confused with herpes simplex, dermatitis herpetiformis and impetigo, and skin reactions caused by contact dermatitis, candidiasis, certain drugs and insect bites.

A number of vaccines against canine distemper exist for dogs (ATCvet code: and combinations) and domestic ferrets (), which in many jurisdictions are mandatory for pets. Infected animals should be quarantined from other dogs for several months owing to the length of time the animal may shed the virus. The virus is destroyed in the environment by routine cleaning with disinfectants, detergents, or drying. It does not survive in the environment for more than a few hours at room temperature (20–25 °C), but can survive for a few weeks in shady environments at temperatures slightly above freezing. It, along with other labile viruses, can also persist longer in serum and tissue debris.

Despite extensive vaccination in many regions, it remains a major disease of dogs.

To prevent canine distemper, puppies should begin vaccination at six to eight weeks of age and then continue getting the “booster shot” every two to four weeks until they are 16 weeks of age. Without the full series of shots, the vaccination will not provide protection against the virus. Since puppies are typically sold at the age of eight to ten weeks, they typically receive the first shot while still with their breeder, but the new owner often does not finish the series. These dogs are not protected against the virus and so are susceptible to canine distemper infection, continuing the downward spiral that leads to outbreaks throughout the country.

Diagnosis relies on viral isolation from tissues, or serological testing with an ELISA. Other methods of diagnosis include Nucleic Acid Testing (NAT), cell culture, and IgM antibody assays. As of September 2016, the Kenya Medical Research Institute (KEMRI) has developed a product called Immunoline, designed to diagnose the disease in humans much faster than in previous methods.

Prophylaxis by vaccination, as well as preventive measures like protective clothing, tick control, and mosquito control are advised. The vaccine for KFDV consists of formalin-inactivated KFDV. The vaccine has a 62.4% effectiveness rate for individuals who receive two doses. For individuals who receive an additional dose, the effectiveness increases to 82.9%. Specific treatments are not available.

Despite decades of research, no vaccines currently exist.

Recombinant technology has however been used to target the formation of vaccines for HPIV-1, -2 and -3 and has taken the form of several live-attenuated intranasal vaccines. Two vaccines in particular were found to be immunogenic and well tolerated against HPIV-3 in phase I trials. HPIV-1 and -2 vaccine candidates remain less advanced.

Vaccine techniques which have been used against HPIVs are not limited to intranasal forms, but also viruses attenuated by cold passage, host range attenuation, chimeric construct vaccines and also introducing mutations with the help of reverse genetics to achieve attenuation.

Maternal antibodies may offer some degree of protection against HPIVs during the early stages of life via the colostrum in breast milk.

If someone is suspected of having polioencephalitis a sample of throat secretions, stool or cerebrospinal fluid is checked for the virus. Blood tests can be done to detect antibodies against viral antigens and foreign proteins. Virus isolation is the most sensitive method and it is most likely to be isolated from stool samples. Once isolated, RT-PCR is used to differentiate naturally occurring strains from vaccine-like strains.

A vaccine has been conditionally approved for use in animals in the US. It has been shown that knockout of the NSs and NSm nonstructural proteins of this virus produces an effective vaccine in sheep as well.

Diagnosis of the oropouche infection is done through classic and molecular virology techniques. These include:

1. Virus isolation attempt in new born mice and cell culture (Vero Cells)

2. Serological assay methods, such as HI (hemagglutination inhibition), NT (neutralization test), and CF (complement fixation test) tests and in-house-enzyme linked immunosorbent assay for total immunoglobulin, IgM, and IgG detection using convalescent sera (this obtained from recovered patients and is rich in antibodies against the infectious agent)

3. Reverse transcription polymerase chain reaction (RT-PCR) and real time RT-PCR for genome detection in acute samples (sera, blood, and viscera of infected animals)

Clinical diagnosis of oropouche fever is hard to perform due to the nonspecific nature of the disease, in many causes it can be confused with dengue fever or other arbovirus illness.

The virus’s transmission cycle in the wild is similar to the continuous sylvatic cycle of yellow fever and is believed to involve wild primates (monkeys) as the reservoir and the tree-canopy-dwelling "Haemagogus" species mosquito as the vector. Human infections are strongly associated with exposure to humid tropical forest environments. Chikungunya virus is closely related, producing a nearly indistinguishable, highly debilitating arthralgic disease. On February 19, 2011, a Portuguese-language news source reported on a recent survey which revealed Mayaro virus activity in Manaus, Amazonas State, Brazil. The survey studied blood samples from 600 residents of Manaus who had experienced a high fever; Mayaro virus was identified in 33 cases. Four of the cases experienced mild hemorrhagic (bleeding) symptoms, which had not previously been described in Mayaro virus disease. The report stated that this outbreak is the first detected in a metropolitan setting, and expressed concern that the disease might be adapting to urban species of mosquito vectors, which would make it a risk for spreading within the country. A study published in 1991 demonstrated that a colonized strain of Brazilian "Aedes albopictus" was capable of acquiring MAYV from infected hamsters and subsequently transmitting it and a study published in October 2011 demonstrated that "Aedes aegypti" can transmit MAYV, supporting the possibility of wider transmission of Mayaro virus disease in urban settings.

Infection with Japanese encephalitis confers lifelong immunity. There are currently three vaccines available: SA14-14-2, IC51 (marketed in Australia and New Zealand as JESPECT and elsewhere as IXIARO) and ChimeriVax-JE (marketed as IMOJEV). All current vaccines are based on the genotype III virus.

A formalin-inactivated mouse-brain derived vaccine was first produced in Japan in the 1930s and was validated for use in Taiwan in the 1960s and in Thailand in the 1980s. The widespread use of vaccine and urbanization has led to control of the disease in Japan, Korea, Taiwan, and Singapore. The high cost of this vaccine, which is grown in live mice, means that poorer countries have not been able to afford to give it as part of a routine immunization program.

The most common adverse effects are redness and pain at the injection site. Uncommonly, an urticarial reaction can develop about four days after injection. Vaccines produced from mouse brain have a risk of autoimmune neurological complications of around 1 per million vaccinations. However where the vaccine is not produced in mouse brains but in vitro using cell culture there is little adverse effects compared to placebo, the main side effects are headache and myalgia.

The neutralizing antibody persists in the circulation for at least two to three years, and perhaps longer. The total duration of protection is unknown, but because there is no firm evidence for protection beyond three years, boosters are recommended every three years for people who remain at risk. Furthermore, there is also no data available regarding the interchangeability of other JE vaccines and IXIARO.

In September 2012 the Indian firm Biological E. Limited has launched an inactivated cell culture derived vaccine based on SA 14-14-2 strain which was developed in a technology transfer agreement with Intercell and is a thiomersal-free vaccine.