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.

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.

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.

A vaccine is available in the UK and Europe, however in laboratory tests it is not possible to distinguish between antibodies produced as a result of vaccination and those produced in response to infection with the virus. Management also plays an important part in the prevention of EVA.

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.

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

Although no specific treatment for acute infection with SuHV1 is available, vaccination can alleviate clinical signs in pigs of certain ages. Typically, mass vaccination of all pigs on the farm with a modified live virus vaccine is recommended. Intranasal vaccination of sows and neonatal piglets one to seven days old, followed by intramuscular (IM) vaccination of all other swine on the premises, helps reduce viral shedding and improve survival. The modified live virus replicates at the site of injection and in regional lymph nodes. Vaccine virus is shed in such low levels, mucous transmission to other animals is minimal. In gene-deleted vaccines, the thymidine kinase gene has also been deleted; thus, the virus cannot infect and replicate in neurons. Breeding herds are recommended to be vaccinated quarterly, and finisher pigs should be vaccinated after levels of maternal antibody decrease. Regular vaccination results in excellent control of the disease. Concurrent antibiotic therapy via feed and IM injection is recommended for controlling secondary bacterial pathogens.

Because of the variability of symptoms, diagnosis is by laboratory testing. Blood samples, nasal swabs and semen can be used for isolation of the virus, detection of the viral RNA by polymerase chain reaction (PCR), and detection of antibodies by ELISA and virus neutralisation tests.

Lesions of paravaccinia virus will clear up with little to no scaring after 4 to 8 weeks. An antibiotic may be prescribed by a physician to help prevent bacterial infection of the lesion area. In rare cases, surgical removal of the lesions can be done to help increase rate of healing, and help minimize risk of bacterial or fungal infection. Upon healing, no long term side effects have been reported.

The CDC recommends screening some pregnant women even if they do not have symptoms of infection. Pregnant women who have traveled to affected areas should be tested between two and twelve weeks after their return from travel. Due to the difficulties with ordering and interpreting tests for Zika virus, the CDC also recommends that healthcare providers contact their local health department for assistance. For women living in affected areas, the CDC has recommended testing at the first prenatal visit with a doctor as well as in the mid-second trimester, though this may be adjusted based on local resources and the local burden of Zika virus. Additional testing should be done if there are any signs of Zika virus disease. Women with positive test results for Zika virus infection should have their fetus monitored by ultrasound every three to four weeks to monitor fetal anatomy and growth.

Diagnosis of paravaccinia virus will often come from Polymerase chain reaction screening ordered by their physician. However, due to how common paravaccinia virus is in rural areas, individuals typically do not seek professional help in diagnosis. Instead individuals may refer to people with local knowledge of the cattle in their area such as ranchers, or veterinarians who have some familiarity with the infections in the region.

A number of various diseases may present with symptoms similar to those caused by a clinical West Nile virus infection. Those causing neuroinvasive disease symptoms include the enterovirus infection and bacterial meningitis. Accounting for differential diagnoses is a crucial step in the definitive diagnosis of WNV infection. Consideration of a differential diagnosis is required when a patient presents with unexplained febrile illness, extreme headache, encephalitis or meningitis. Diagnostic and serologic laboratory testing using polymerase chain reaction (PCR) testing and viral culture of CSF to identify the specific pathogen causing the symptoms, is the only currently available means of differentiating between causes of encephalitis and meningitis.

However, simple husbandry changes and practical midge control measures may help break the livestock infection cycle. Housing livestock during times of maximum midge activity (from dusk to dawn) may lead to significantly reduced biting rates. Similarly, protecting livestock shelters with fine mesh netting or coarser material impregnated with insecticide will reduce contact with the midges. The "Culicoides" midges that carry the virus usually breed on animal dung and moist soils, either bare or covered in short grass. Identifying breeding grounds and breaking the breeding cycle will significantly reduce the local midge population. Turning off taps, mending leaks and filling in or draining damp areas will also help dry up breeding sites. Control by trapping midges and removing their breeding grounds may reduce vector numbers. Dung heaps or slurry pits should be covered or removed, and their perimeters (where most larvae are found) regularly scraped.

Preliminary diagnosis is often based on the patient's clinical symptoms, places and dates of travel (if patient is from a nonendemic country or area), activities, and epidemiologic history of the location where infection occurred. A recent history of mosquito bites and an acute febrile illness associated with neurologic signs and symptoms should cause clinical suspicion of WNV.

Diagnosis of West Nile virus infections is generally accomplished by serologic testing of blood serum or cerebrospinal fluid (CSF), which is obtained via a lumbar puncture. Initial screening could be done using the ELISA technique detecting immunoglobulins in the sera of the tested individuals.

Typical findings of WNV infection include lymphocytic pleocytosis, elevated protein level, reference glucose and lactic acid levels, and no erythrocytes.

Definitive diagnosis of WNV is obtained through detection of virus-specific antibody IgM and neutralizing antibodies. Cases of West Nile virus meningitis and encephalitis that have been serologically confirmed produce similar degrees of CSF pleocytosis and are often associated with substantial CSF neutrophilia.

Specimens collected within eight days following onset of illness may not test positive for West Nile IgM, and testing should be repeated. A positive test for West Nile IgG in the absence of a positive West Nile IgM is indicative of a previous flavavirus infection and is not by itself evidence of an acute West Nile virus infection.

If cases of suspected West Nile virus infection, sera should be collected on both the acute and

convalescent phases of the illness. Convalescent specimens should be collected 2–3 weeks after acute specimens.

It is common in serologic testing for cross-reactions to occur among flaviviruses such as dengue virus (DENV) and tick-borne encephalitis virus; this necessitates caution when evaluating serologic results of flaviviral infections.

Four FDA-cleared WNV IgM ELISA kits are commercially available from different manufacturers in the U.S., each of these kits is indicated for use on serum to aid in the presumptive laboratory diagnosis of WNV infection in patients with clinical symptoms of meningitis or encephalitis. Positive WNV test results obtained via use of these kits should be confirmed by additional testing at a state health department laboratory or CDC.

In fatal cases, nucleic acid amplification, histopathology with immunohistochemistry, and virus culture of autopsy tissues can also be useful. Only a few state laboratories or other specialized laboratories, including those at CDC, are capable of doing this specialized testing.

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.

SuHV1 can be used to analyze neural circuits in the central nervous system (CNS). For this purpose the attenuated (less virulent) Bartha SuHV1 strain is commonly used and is employed as a retrograde and anterograde transneuronal tracer. In the retrograde direction, SuHV1-Bartha is transported to a neuronal cell body via its axon, where it is replicated and dispersed throughout the cytoplasm and the dendritic tree. SuHV1-Bartha released at the synapse is able to cross the synapse to infect the axon terminals of synaptically connected neurons, thereby propagating the virus; however, the extent to which non-synaptic transneuronal transport may also occur is uncertain. Using temporal studies and/or genetically engineered strains of SuHV1-Bartha, second, third, and higher order neurons may be identified in the neural network of interest.

A blood test is the only way to confirm a case of Ross River Fever. Several types of blood tests may be used to examine antibody levels in the blood. Tests may either look for simply elevated antibodies (which indicate some sort of infection), or specific antibodies to the virus.

For infants with suspected congenital Zika virus disease, the CDC recommends testing with both serologic and molecular assays such as RT-PCR, IgM ELISA and plaque reduction neutralization test (PRNT). RT-PCR of the infants serum and urine should be performed in the first two days of life. Newborns with a mother who was potentially exposed and who have positive blood tests, microcephaly or intracranial calcifications should have further testing including a thorough physical investigation for neurologic abnormalities, dysmorphic features, splenomegaly, hepatomegaly, and rash or other skin lesions. Other recommended tests are cranial ultrasound, hearing evaluation, and eye examination. Testing should be done for any abnormalities encountered as well as for other congenital infections such as syphilis, toxoplasmosis, rubella, cytomegalovirus infection, lymphocytic choriomeningitis virus infection, and herpes simplex virus. Some tests should be repeated up to 6 months later as there can be delayed effects, particularly with hearing.

A recent study from The Cleveland Clinic reported that BK viremia load > 185 000 copies/ml at the time of first positive BKV diagnosis - to be the strongest predictor for BKVAN (97% specificity and 75% sensitivity). In addition the BKV peak viral loads in blood reaching 223 000 copies/ml at any time was found to be predictive for BKVAN (91% specificity and 88% sensitivity) .

Currently, there is no proven, safe treatment for monkeypox. The people who have been infected can be vaccinated up to 14 days after exposure.

A range of laboratory investigations are performed, where possible, to diagnose the disease and assess its course and complications. The confidence of a diagnosis can be compromised by if laboratory tests are not available. One comprising factor is the number of febrile illnesses present in Africa, such as malaria or typhoid fever that could potentially exhibit similar symptoms, particularly for non-specific manifestations of Lassa fever. In cases with abdominal pain, in countries where Lassa is common, Lassa fever is often misdiagnosed as appendicitis and intussusception which delays treatment with the antiviral ribavirin. In West Africa, where Lassa is most prevalent, it is difficult for doctors to diagnose due to the absence of proper equipment to perform tests.

The FDA has yet to approve a widely validated laboratory test for Lassa, but there are tests that have been able to provide definitive proof of the presence of the LASV virus. These tests include cell cultures, PCR, ELISA antigen assays, plaque neutralization assays, and immunofluorescence essays. However, immunofluorescence essays provide less definitive proof of Lassa infection. An ELISA test for antigen and IgM antibodies give 88% sensitivity and 90% specificity for the presence of the infection. Other laboratory findings in Lassa fever include lymphopenia (low white blood cell count), thrombocytopenia (low platelets), and elevated aspartate aminotransferase levels in the blood. Lassa fever virus can also be found in cerebrospinal fluid.

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.

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.