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.

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.

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.

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.

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.

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

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.

The differential diagnosis in a case of suspected human rabies may initially include any cause of encephalitis, in particular infection with viruses such as herpesviruses, enteroviruses, and arboviruses such as West Nile virus. The most important viruses to rule out are herpes simplex virus type one, varicella zoster virus, and (less commonly) enteroviruses, including coxsackieviruses, echoviruses, polioviruses, and human enteroviruses 68 to 71.

New causes of viral encephalitis are also possible, as was evidenced by the 1999 outbreak in Malaysia of 300 cases of encephalitis with a mortality rate of 40% caused by Nipah virus, a newly recognized paramyxovirus. Likewise, well-known viruses may be introduced into new locales, as is illustrated by the outbreak of encephalitis due to West Nile virus in the eastern United States. Epidemiologic factors, such as season, geographic location, and the patient's age, travel history, and possible exposure to bites, rodents, and ticks, may help direct the diagnosis.

A Zika virus infection might be suspected if symptoms are present and an individual has traveled to an area with known Zika virus transmission. Zika virus can only be confirmed by a laboratory test of body fluids, such as urine or saliva, or by blood test.

, no approved vaccines are available. A phase-II vaccine trial used a live, attenuated virus, to develop viral resistance in 98% of those tested after 28 days and 85% still showed resistance after one year. However, 8% of people reported transient joint pain, and attenuation was found to be due to only two mutations in the E2 glycoprotein. Alternative vaccine strategies have been developed, and show efficacy in mouse models. In August 2014 researchers at the National Institute of Allergy and Infectious Diseases in the USA were testing an experimental vaccine which uses virus-like particles (VLPs) instead of attenuated virus. All the 25 people participated in this phase 1 trial developed strong immune responses. As of 2015, a phase 2 trial was planned, using 400 adults aged 18 to 60 and to take place at 6 locations in the Caribbean. Even with a vaccine, mosquito population control and bite prevention will be necessary to control chikungunya disease.

Laboratory blood tests can identify evidence of chikungunya or other similar viruses such as dengue and Zika. Blood test may confirm the presence of IgM and IgG anti-chikungunya antibodies. IgM antibodies are highest 3 to 5 weeks after the beginning of symptoms and will continue be present for about 2 months.

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.

Viral entry is the earliest stage of infection in the viral life cycle, as the virus comes into contact with the host cell and introduces viral material into the cell. The major steps involved in viral entry are shown below. Despite the variation among viruses, there are several shared generalities concerning viral entry.

Vaccines are available (ATCvet codes: for the inactivated vaccine, for the live vaccine, plus various combinations).

Given that avian reovirus infections are widespread, the viruses are relatively resistant outside the host, and that vertical and horizontal transmission occurs, eradicating avian reovirus infection in commercial chicken flocks is very unlikely. In addition, absence of detectable seroconversion and failure to detect virus in cloacal swabs are unreliable indicators of resisting infection, or transmission via the egg. Thus, the most proactive and successful approach to controlling this disease is through vaccination. Since chicks are more prone to being detrimentally affected by the disease right after hatching, vaccine protocols that use live and killed vaccines are designed to provide protection during the very early stages of life. This approach has been accomplished through active immunity after early vaccination and a live vaccine or passive immunity from maternal antibodies followed with vaccination of the breeder hens. Currently, efforts toward administering inactivated or live vaccines to breeding stock to allow passive immunity to the offspring via the yolk are being taken.

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.

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.

Early symptoms of EVD may be similar to those of other diseases common in Africa, including malaria and dengue fever. The symptoms are also similar to those of other viral hemorrhagic fevers such as Marburg virus disease.

The complete differential diagnosis is extensive and requires consideration of many other infectious diseases such as typhoid fever, shigellosis, rickettsial diseases, cholera, sepsis, borreliosis, EHEC enteritis, leptospirosis, scrub typhus, plague, Q fever, candidiasis, histoplasmosis, trypanosomiasis, visceral leishmaniasis, measles, and viral hepatitis among others.

Non-infectious diseases that may result in symptoms similar to those of EVD include acute promyelocytic leukemia, hemolytic uremic syndrome, snake envenomation, clotting factor deficiencies/platelet disorders, thrombotic thrombocytopenic purpura, hereditary hemorrhagic telangiectasia, Kawasaki disease, and warfarin poisoning.

Rabies can be difficult to diagnose, because, in the early stages, it is easily confused with other diseases or with aggressiveness. The reference method for diagnosing rabies is the fluorescent antibody test (FAT), an immunohistochemistry procedure, which is recommended by the World Health Organization (WHO). The FAT relies on the ability of a detector molecule (usually fluorescein isothiocyanate) coupled with a rabies-specific antibody, forming a conjugate, to bind to and allow the visualisation of rabies antigen using fluorescent microscopy techniques. Microscopic analysis of samples is the only direct method that allows for the identification of rabies virus-specific antigen in a short time and at a reduced cost, irrespective of geographical origin and status of the host. It has to be regarded as the first step in diagnostic procedures for all laboratories. Autolysed samples can, however, reduce the sensitivity and specificity of the FAT. The RT PCR assays proved to be a sensitive and specific tool for routine diagnostic purposes, particularly in decomposed samples or archival specimens. The diagnosis can be reliably made from brain samples taken after death. The diagnosis can also be made from saliva, urine, and cerebrospinal fluid samples, but this is not as sensitive and reliable as brain samples. Cerebral inclusion bodies called Negri bodies are 100% diagnostic for rabies infection but are found in only about 80% of cases. If possible, the animal from which the bite was received should also be examined for rabies.

Some light microscopy techniques may also be used to diagnose rabies at a tenth of the cost of traditional fluorescence microscopy techniques, allowing identification of the disease in less-developed countries.

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.

Possible non-specific laboratory indicators of EVD include a low platelet count; an initially decreased white blood cell count followed by an increased white blood cell count; elevated levels of the liver enzymes alanine aminotransferase (ALT) and aspartate aminotransferase (AST); and abnormalities in blood clotting often consistent with disseminated intravascular coagulation (DIC) such as a prolonged prothrombin time, partial thromboplastin time, and bleeding time. Filovirions, such as EBOV, may be identified by their unique filamentous shapes in cell cultures examined with electron microscopy, but this method cannot distinguish the various filoviruses.

The specific diagnosis of EVD is confirmed by isolating the virus, detecting its RNA or proteins, or detecting antibodies against the virus in a person's blood. Isolating the virus by cell culture, detecting the viral RNA by polymerase chain reaction (PCR) and detecting proteins by enzyme-linked immunosorbent assay (ELISA) are methods best used in the early stages of the disease and also for detecting the virus in human remains. Detecting antibodies against the virus is most reliable in the later stages of the disease and in those who recover. IgM antibodies are detectable two days after symptom onset and IgG antibodies can be detected 6 to 18 days after symptom onset. During an outbreak, isolation of the virus via cell culture methods is often not feasible. In field or mobile hospitals, the most common and sensitive diagnostic methods are real-time PCR and ELISA. In 2014, with new mobile testing facilities deployed in parts of Liberia, test results were obtained 3–5 hours after sample submission. In 2015 a rapid antigen test which gives results in 15 minutes was approved for use by WHO. It is able to confirm Ebola in 92% of those affected and rule it out in 85% of those not affected.

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.

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.

Definitive diagnosis is usually made at a reference laboratory with advanced biocontainment capabilities. The findings of laboratory investigation vary somewhat between the viruses but in general there is a decrease in the total white cell count (particularly the lymphocytes), a decrease in the platelet count, an increase in the blood serum liver enzymes, and reduced blood clotting ability measured as an increase in both the prothrombin (PT) and activated partial thromboplastin times (PTT). The hematocrit may be elevated. The serum urea and creatine may be raised but this is dependent on the hydration status of the patient. The bleeding time tends to be prolonged.