Chapter 2. Laboratory Diagnosis of Viral Diseases

Viruses remain a continuing threat to humankind regardless of age, gender, ethnicity, and socioeconomic status. They are a leading cause of morbidity and mortality worldwide, and severe illness commonly occurs in infants, the elderly, the chronically ill, the malnourished, and the immunocompromised.

The clinical diagnosis of viral diseases can be difficult, and laboratory confirmation is required in patients with serious illnesses since signs and symptoms are often overlapping and not always specific for any one viral agent. This is particularly true in children. Early detection of viral diseases can have a prompt and significant impact on patient care by increasing clinical awareness and providing for more informed decision making for better patient management.1–8 A rapid and specific diagnosis can help in decreasing the use of unwarranted laboratory tests, hospital procedures, and antimicrobial drugs, resulting in reduced costs related to supportive care and a reduction in hospital stay.1,3,4,8–13 In some cases where antiviral therapy is available, rapid and accurate viral detection can provide the opportunity for prophylaxis against certain viral infections or early treatment that may limit the extent of disease and reduce associated sequelae.1,14 Finally, timely laboratory surveillance for viruses can provide for rapid outbreak identification and assist in the prevention of community and hospital spread.15–18

Many methods are available for the diagnosis and management of viral diseases (Table 2–1).19,20 These include (1) cell culture systems for the isolation of viruses in human or animal cells, (2) rapid immunologic and molecular assays for the direct detection and/or quantification of viral proteins or nucleic acids, (3) serologic tests to detect virus-specific IgG and/or IgM antibody responses, (4) electron microscopy to identify viruses based on size and shape of viral particles, (5) histologic and cytologic techniques for detection of viral-induced morphological changes within tissues and exfoliated cells, and (6) genotypic and phenotypic assays to detect antiviral drug resistance and to identify genetic variants that may not respond to therapy. There are notable differences in the use and clinical performance of these methods and the relative importance of certain tests has changed over the years. Therefore, the selection of which assays to perform and the choice of specimens to collect for testing should be made judiciously and in consultation with appropriate laboratory personnel and will depend on the patient population and clinical situation, the intended use of the individual tests, and the capabilities and resources of individual laboratories.

Appropriate specimen collection and handling is absolutely critical to the success of laboratory testing for diagnosis of viral infections.21 Irrespective of the location of testing or techniques used, specimens that are poorly collected, ill timed, or incorrectly handled between the time of collection and testing are more likely to yield false test results. A comprehensive listing of the selection of viral specimens based on clinical infections, the suspected viral agents, and the tests to be performed are listed in Tables 2–2, 2–3, 2–4, 2–5, 2–6, 2–7, 2–8, 2–9, 2–10, and 2–11.

In general, specimens should be collected as close to clinical onset as possible (e.g., within the first 1–3 days). Viral shedding is at its maximum at this time and increases the likelihood of virus detection. Most acute viral infections are self-limiting and last for approximately 5–10 days, so delays in specimen collection may adversely affect laboratory testing. The amount of virus present and the duration of viral shedding will vary, however, and depends on the age of the patient, the virus, a competent immune system, the anatomical site chosen for specimen collection, and whether the infection is localized or systemic. Young infants and immunosuppressed children may shed virus for more extended times. The specimen site should be determined by the clinical presentation and the pathogenic potential of the suspected virus (Tables 2–2, 2–3, 2–4, 2–5, 2–6, 2–7, 2–8, 2–9, 2–10, and 2–11). Recovery of a given virus may be enhanced by collecting the same specimen type over the course of several days or by collecting various specimens from different body sites. Specimens should be transported to the laboratory as quickly as possible after collection. This is especially true when attempting to grow viruses from clinical specimens since some viruses, particularly those with envelopes, are quite labile outside their natural host and do not survive well under adverse environmental conditions. When immediate transport is not possible, refrigerate the specimens or keep them on wet ice. For delays of more than 24–48 hours, specimens should be processed as needed and rapidly frozen to −70°C and then transported to the laboratory on dry ice. Specimens to be used for the direct detection of viruses (e.g., culture, antigen assays, or molecular methods) should never be stored at room temperature or frozen at −20°C; it is acceptable to freeze serum or plasma at −20°C for transport to and extended storage in the laboratory when being used for detection of viral-specific antibodies. It is recommended that all specimens for molecular testing be stored at 4°C immediately after collection and then promptly sent to the laboratory for appropriate processing and storage for testing. This is critical to ensure the stability and amplification of nucleic acids, particularly for the detection of RNA viruses since RNA is unstable and easily degraded by RNases from the surrounding environment.

Collection Instructions for Selected Specimen Types

Blood for Plasma or White Blood Cells

Approximately 4–7 mL of whole blood should be collected in a suitable anticoagulant such as ethylenediamminetriaminoacetic acid (EDTA), sodium heparin, sodium citrate, or acid citrate dextrose. EDTA is currently the preferred anticoagulant for most viral studies that require plasma or white blood cells and is considered to be the best stabilizer of nucleic acids for molecular testing.

Blood for Serum

Serum is the preferred specimen for most serological assays. Approximately 1–2 mL of blood should be collected for every two to three tests ordered; collection tubes should not contain anticoagulants or preservatives. A single serum specimen is sufficient to determine the immune status of an individual or for the detection of virus-specific IgM antibody. For the diagnosis of current or recent viral infections, paired serum specimens collected 10–14 days apart are needed when testing for virus-specific IgG antibody. There are exceptions to this general rule and only a single serum for IgG is required for viruses such as Epstein–Barr virus (EBV), hepatitis B virus (HBV), and parvovirus B19. The acute-phase serum should be collected as early as possible in the course of the illness, and the acute and convalescent sera should be tested simultaneously to obtain the most meaningful results. Serum specimens from mother, fetus, and newborn can be submitted for the evaluation of IgG and IgM antibodies for the detection of prenatal, natal, and postnatal viral infections with cytomegalovirus (CMV), rubella virus, herpes simplex virus (HSV), varicella-zoster virus (VZV), parvovirus B19, human immunodeficiency virus (HIV), HBV, hepatitis C virus (HCV), and others. Plasma can be used as an acceptable alternative to serum for serological diagnosis of HIV and hepatitis viruses. Oral mucosal transudates rich in gingival crevicular fluid,22,24 urine,23 and dried blood spots24,25 can be used as noninvasive alternatives to the collection of blood for the detection of HIV-specific antibodies. Cerebrospinal fluid (CSF) may be collected and paired with a serum specimen from the same date to be tested for viral-specific antibody in patients with viral neurological diseases.26 However, the yield of such testing is low for most viruses and limited by delays in intrathecal production of antibody, and many hospital laboratories no longer perform antibody testing on CSF samples. Testing for viral-specific antibodies in CSF may be beneficial for certain viruses, including the arboviruses, measles, mumps, and rabies viruses, herpes B virus, and lymphocytic choriomeningitis virus (LCMV).

Respiratory Specimens

Traditionally, nasopharyngeal aspirates or nasopharyngeal washes have been the preferred specimens for the detection of respiratory viruses, and are felt to be superior to swabs for the adequate collection of respiratory epithelial cells. Aspirates are particularly useful for infants and young children who produce copious amounts of mucus during their viral respiratory illness, but are not as practical in older children and adults with less respiratory secretions to aspirate. In the latter populations, swabs are far more convenient for medical personnel to use and patients are more willing to allow collection of this specimen type. For optimum recovery of viruses from the respiratory tract when using swabs, collection of combined throat and nasopharyngeal swabs is recommended since recovery of certain viruses can vary depending on the amount of virus present in either the throat or nose. For instance, nasopharyngeal swabs are better than throat swabs for the recovery of seasonal human influenza virus strains, while throat swabs are preferred in humans with respiratory disease caused by avian H5N1 influenza virus since the throat contains higher titers of this virus. Tracheal or transtracheal aspirates or bronchoalveolar lavage specimens are superior for the indication of lower respiratory tract infections. Sputum has been used for the detection of respiratory viruses, but normally contains less than the optimum amount of viruses required for testing and is not recommended. Regardless of the respiratory specimen chosen for collection, children generally shed respiratory viruses at higher titers for longer periods than adults. Excess dilution of respiratory specimens during collection may decrease test performance in the laboratory and should be avoided.

Nasopharyngeal Aspirate

Obtain a mucus trap, a sterile French gauge suction catheter, and a vacuum aspiration pump. The length and diameter of the catheter and suction pressure to be used will vary with the age of the patient and should be appropriate for an infant, child, or adult. Attach the catheter to the trap and then connect the trap to the vacuum aspiration pump. Insert the catheter into the nose, directed posteriorly and toward the opening of the external ear. The depth of the catheter insertion necessary to reach the posterior pharynx is equivalent to the distance between the anterior nares and external opening of the ear. Apply vacuum aspiration. Using a rotating movement, collect as much of the secretions as possible and then slowly withdraw the catheter. The catheter should remain in the nasopharynx for no longer than 10 seconds. Hold the trap upright to prevent the secretions from going into the pump. Rinse the catheter (if necessary) with approximately 2 mL of viral transport medium if the secretions do not move freely through the tubing into the trap; disconnect the suction catheter and vacuum pump and close off the trap for transport to the laboratory.

Nasal Wash: Syringe Method

Obtain a 3–5-cc syringe and fill the syringe with sterile saline. Attach an appropriately sized French gauge catheter to the syringe, and, as above, insert the tubing into the nose to reach the posterior nasopharynx. Quickly instill the saline into the nasopharynx and immediately withdraw the fluid back into the syringe to recover the specimen material. Inject the aspirated specimen from the syringe into a suitable dry, sterile container or one containing a specified volume of viral transport medium.

Nasal Wash: Bulb Method

Suction 3–5 mL of saline into a clean sterile 1–2 oz. tapered rubber bulb. Insert the bulb into one nostril until the nostril is occluded. Instill the saline into the nostril with one squeeze of the bulb and immediately release the bulb to recover the nasal specimen. Empty the bulb into a suitable dry, sterile container or one containing a specified volume of viral transport medium.

Nasopharyngeal or Oropharyngeal Swabs

(See below for swab specimens.)

Sterile Body Fluids

Urine, CSF, amniotic, pleural and pericardial fluids, and other sterile body fluid specimens should be submitted to the laboratory in sterile, leak proof containers. Do not dilute these specimens in viral transport medium. All body fluid specimens must be of sufficient quantity, and, as a general rule, should equal a minimum of 0.5 mL per test requested. Shedding of some viruses in urine may be intermittent (e.g., CMV) and collecting two to three urine specimens over time may enhance recovery. Sampling of amniotic fluid should be done after 21–23 weeks of fetal gestation for best testing sensitivity when using either serological or direct detection methods for prenatal diagnosis.

Stool

Stool is the specimen of choice for the detection of enteric viruses. Approximately 2–4 mL of liquid stool or 2–4 g of formed stool should be placed in leak proof containers that do not contain media, preservatives, serum, or detergent, as any of these additives may interfere with specific antigen tests for rotavirus and adenovirus types 40 and 41.

Swab Specimens

Swabs are used for collecting specimens from dermal, rectal, respiratory, and ocular sites. Plastic- or metal-shafted swabs with rayon, Dacron, cotton, or polyester tips should be used; calcium alginate or wooden-shafted swabs are inhibitory to some viruses and are not acceptable. To facilitate binding of cellular material to the swab, the swab can be premoistened with sterile saline. The excess liquid should be adequately expressed from the swab before attempting to collect the specimen. All swab specimens should be placed in viral transport medium immediately after collection. Recently, a new type of swab made with flocked nylon has been developed by Copan Diagnostics (Corona, CA). Use of this swab has resulted in enhanced specimen recovery and improved test performance by allowing for increased absorption of cells to the swab during collection and improved release of the cells from the swab during processing and testing in the laboratory.27,28

Eye (Conjunctivae)

Conjunctival swabs should be taken by stroking the lower conjunctival sac of the eye five to six times with a sterile swab.

Dermal Lesions

Cells and fluid from fresh vesicles are superior to specimens prepared from other lesion types (e.g., vesicles > pustules > ulcers > crusts) for direct antigen detection, polymerase chain reaction (PCR), and viral culture. Unroof the vesicle with a sterile needle or scalpel blade and use a sterile swab to vigorously swab the margins and base of the lesion to obtain infected epithelial cells. For ulcers, use a swab to remove pus without disrupting the lesion base and then use a fresh sterile swab to firmly swab the lesion as described above. Crusted lesions should have the crust removed and discarded before collection of cells from the lesion base and margins. The yield of HSV or VZV from a crusted lesion is low for rapid antigen and culture-based tests and moderate for PCR, so the utility of collecting specimens from this type of lesions should be considered.

Nasopharyngeal

Insert a swab into the posterior nasopharynx. Press the swab tip on the mucosal surface of the midinferior portion of the inferior turbinate and rub or rotate the swab tip several times across the mucosal surface to loosen and collect respiratory epithelial cells.

Oropharyngeal

Gently open the mouth or ask the patient (if age appropriate) to open the mouth widely and phonate an “ah” to lift the uvula and reduce the gag reflex. Gently depress the tongue with a tongue blade and guide a swab into the posterior oropharynx. Use a gentle but firm back-and-forth sweeping motion to swab thoroughly behind the uvula and between the tonsillar pillars while avoiding the buccal mucosa, tonsils, tongue, and palate. Collected nasopharyngeal and oropharyngeal swabs should be combined and placed into a single tube of viral transport medium. Doing so usually improves on the number of epithelial cells available for testing and enhances the recovery of viruses from these sources.

Rectal

Rectal swabs are normally inferior to stool specimens for the detection of enteric viruses and their collection is discouraged. If it is necessary to collect a rectal swab, a sufficient quantity of fecal material (e.g., at least a pea-sized amount) should be obtained. If fecal material is not clearly visible on the swab, the specimen is most likely inadequate. A swab of the rectal mucosa can be performed if viral proctitis is suspected.

Tissues

Freshly collected tissue specimens should be immediately placed in viral transport medium to prevent drying.

All specimens submitted to the laboratory for diagnostic testing should be labeled with the patient's full name, the medical record number or other unique identifier, date and time of collection, and signature of the collector. Each specimen should be accompanied by a requisition slip containing the same information as on the specimen container as well as the patient's location within the hospital, physician's name, and contact information, specimen site, test(s) requested, virus(es) suspected, and any other relevant clinical data (e.g., clinical diagnosis, current therapy, or factors affecting immune status such as transplantation or malignancy). Specimens must be appropriate for the test(s) requested and/or the clinical situation described. Meaningful communication between health care providers and laboratorians is especially important to determine the appropriateness of specimens for viral diagnosis.

See Tables 2–2, 2–3, 2–4, 2–5, 2–6, 2–7, 2–8, 2–9, 2–10, and 2–11 for selecting the most appropriate laboratory tests depending on the clinical situation and suspected virus or viruses.

Cell Culture Systems

The isolation and identification of viruses in cell culture remains an important and integral component of most clinical virology laboratories29–30, and advances in technology over the years have improved the speed and usefulness of cell culture systems for the detection of certain viruses.

Conventional Tube Cultures

Conventional tube culture systems have long been the cornerstone of diagnostic virology and are the traditional counterpart to growing bacteria in the microbiology laboratory. Unlike bacteria that grow in nutrient broth or on solid agar media, viruses are obligate intracellular parasites and, therefore, require living cells to replicate. Clinical specimens are normally inoculated onto various tissue culture cell lines of animal and human origin grown in 16 × 125-mm glass or plastic tubes and incubated under conditions suitable for isolation of the largest number of viruses. The cultures are examined over a period of time using a standard light microscope, and the presence of viral growth is usually recognized by the development of a virus-induced cytopathic effect (CPE) within infected cells. Each virus that may grow in culture has its characteristic CPE; the rate at which CPE progresses, the type of CPE observed, and the cell line in which the virus replicates are factors in determining the type of virus present. Some viruses, most notably influenza and parainfluenza viruses, often do not produce CPE as primary isolates and their growth in culture is detected by alternative techniques such as measuring hemagglutinin activity and using monoclonal antibodies to visualize specific viral proteins in an immunofluorescence assay. Conventional tube cultures offer the distinct advantage of being able to isolate a wide range of viruses from a given clinical specimen and can detect unknown or unsuspected viral agents. Also, the growth of a virus in a culture is highly specific depending on which cell lines are selected and how the culture system is designed, and low titers of virus present in a specimen can be amplified to sufficient levels for detection and further characterization. Conversely, conventional tube cultures are slow, often taking many days to weeks to obtain a final result and have a limited impact on clinical decision making. They also are time-consuming, labor-intensive, require specialized facilities and expertise, and have a varied sensitivity as many medically relevant viruses are very difficult to grow or cannot be grown at all. Alternatives to conventional tube cultures have been developed and are discussed below. These include shell vial/multiwell plate cultures and the use of genetically engineered and mixed-cell populations.

Shell Vial/Multiwell Plate Cultures

Shell vial or multiwell plate cultures were originally designed to decrease the time required for detection of viruses in culture, especially slow-growing viruses like CMV and VZV. With this method, low-speed centrifugation is used to inoculate clinical specimens onto the surface of cell monolayers grown on 12-mm round coverslips in the bottom of 1-dram (3.7 mL) vials or flat-bottomed 24- or 48-well plates to facilitate viral infection of the cells. Following incubation, viral growth is detected prior to the development of CPE using immunofluorescence staining with monoclonal antibodies directed against specific viral proteins. This method has been applied to the rapid detection of CMV, HSV, VZV, enteroviruses, measles and rubella viruses, and respiratory viruses such as respiratory syncytial virus (RSV), influenza virus types A and B, parainfluenza virus types 1, 2, 3, and 4, adenovirus, and more recently, human metapneumovirus. 31 The major advantage of this culture system over conventional tube cultures is that results are available in a much shorter time frame (most in 24–48 hours, but may take up to 5 days for some respiratory viruses). Disadvantages include that only viruses that are being sought can be identified and only one or a few viruses can be detected at a time. Also, this type of culture is normally less sensitive than conventional tube culture systems, and similar to conventional cultures, involves considerable time, labor, and expertise to perform.

Genetically Engineered Cells

Transgenic cells that have been genetically modified to promote the stable expression of a process or processes required by the virus for entry into or replication within cells have been developed and are being used to facilitate virus detection in culture.32 An enzyme-linked virus-inducible culture system (ELVIS) has been commercially developed by Diagnostic Hybrids (Athens, OH) for the rapid detection of HSV-1 and HSV-2.33,34 The system uses genetically engineered baby hamster kidney cells containing the lac Z gene of Escherichia coli driven by the ICP6 promoter from the UL39 gene of HSV-1. Clinical specimens are inoculated by centrifugation onto cell monolayers in 24-well plates and incubated for 16–24 hours. Cells infected only with HSV express β-galactosidase and are identified by their blue color following histochemical staining for β-galactosidase activity and examination by light microscopy. The assay is rapid and relatively simple to perform and requires less labor and expertise than the more conventional centrifugation-assisted cultures described above. It has the important advantage of easily detecting a color change that is readily induced following infection with either HSV type. To enhance the recovery of enteroviruses in culture, buffalo green monkey kidney (BGMK) cells have been transfected with the gene for human decay-accelerating factor (hDAF), a receptor that certain enterovirus types interact with during entry into a cell.35 The developed genetically engineered cell line, BGMK-hDAF, has an expanded host range and increased sensitivity for the detection of various enteroviruses and has been commercialized for use in a mixed-cell culture system like the one described below. A genetically engineered human embryonic kidney 293T cell line with a reporter gene inducible by influenza A virus has also been developed for the rapid detection of multiple strains of influenza A.36 The major limitations of these transgenic cell lines are that they will detect only the virus for which they were designed and they are available for only a small number of viruses, thereby limiting their diagnostic use.

Mixed-Cell Populations

A more recent development in cell culture involves the mixing of two cell lines together to form a single monolayer in shell vials or multiwell plates. As above, clinical specimens are inoculated by centrifugation onto the monolayers and viruses that can grow in either or both of the mixed cell lines and are detected by immunofluorescence using viral-specific monoclonal antibodies conjugated with different fluorochromes. The major advantage of using mixed cell populations is that multiple viruses can be detected in a single vial or well-thereby decreasing the need to use multiple single tubes as is done with conventional tube cultures. The mixed-cell culture system also takes advantage of the speed involved in virus detection when using the techniques of centrifugation-assisted inoculation and identification of viruses prior to the development of CPE. Cocultivated cell lines of this type are now commercially available for the detection of HSV and VZV,37 enteroviruses,35 and respiratory viruses, which include RSV, influenza virus types A and B, parainfluenza virus types 1, 2, and 3, and adenovirus.38–40

In general, the use and relative importance of cell culture systems for viral isolation is declining with the continued development of rapid and accurate immunologic and molecular tests. However, viral isolation is still one of the most practical and convenient methods for many clinical virology laboratories that are attempting to diagnose viral diseases. Also, specific needs for culture-based systems will most likely always remain. For example, laboratory confirmation of antiviral drug resistance may be important in certain clinical situations involving viruses such as CMV, HSV, VZV, HIV, and influenza A and B viruses. To this end, culture-based, phenotypic antiviral susceptibility assays have been developed (see Ref. 41 for a review). Virus replication within cultured cells is measured in the presence or absence of an antiviral drug and the susceptibility of the virus is expressed as the drug concentration required to inhibit viral replication by 50% relative to infected cell cultures containing no drug. The disadvantages of these tests are that they are relatively costly and labor-involved, and usually have turnaround times of 2–6 weeks depending on the virus examined and the assay used. They are also only available in a limited number of reference laboratories. The major advantage is that culture-based, phenotypic assays are a direct measure of the action of an antiviral drug on a live, growing virus.

Direct Detection Tests

Antigen Detection Assays

Immunologic tests for direct detection of viral antigens in clinical material are now commercially available for many viruses and all or some of the assays are routinely used in most clinical laboratories. The tests are rapid, inexpensive, relatively simple to perform, and, unlike culture-based systems, do not require viable virus for detection. The sensitivity and specificity of these tests are variable and are highly dependent on the virus to be detected, the testing format, and the quality of the specimen. In general, antigen detection assays are usually not as accurate as viral culture or molecular amplification techniques. Also, similar to the centrifugation-assisted cultures described above, direct antigen detection assays can only identify the specific viruses for which the test was designed.

Immunofluorescence

Immunofluorescence is used extensively for the direct visualization of antigens of a number of viruses,42–48 including CMV, EBV, HSV, VZV, RSV, influenza virus types A and B, parainfluenza virus types 1, 2, 3, and 4, adenovirus, metapneumovirus, and measles, mumps, rubella, and rabies viruses. Monoclonal antibodies are now available for these and many other less common viruses, and manufacturers now provide kits that contain all of the necessary reagents for staining. When using immunofluorescence, cells from submitted clinical specimens are fixed to the surface of glass slides and viral antigens within infected cells are detected using virus-specific primary antibodies. A direct or indirect immunofluorescence assay can be used to detect the antigen–antibody complexes. In the direct method, the virus-specific antibody is directly conjugated with a fluorochrome, while in the indirect method, the primary antibody is allowed to react with viral antigen and the specific complexes are detected using an antispecies-specific antibody conjugated with a fluorochrome. The choice of the method to be used depends on the availability and quality of conjugated and unconjugated antibodies and the particular viral antigen to be detected. The assays normally take 1–3 hours to complete and microscopic examination of the slides requires considerable expertise for correct reading and interpretation of the results. By using immunofluorescence, the quality of specimens can be assessed and specimens can be screened for multiple viruses of interest depending on the number of available antibodies. Immunofluorescence is more likely to be performed in larger academic medical centers and public health laboratories rather than in community hospitals because of the need for an immunofluorescence microscope and the requirement for a higher level of expertise to perform these assays. Also, the method is not available for viral infections caused by enteroviruses, rhinoviruses, and viral agents of gastroenteritis because of the limited availability of appropriate reagents. An immunofluorescence test for CMV, called the antigenemia assay, is widely used for the detection and quantification of CMV from blood leukocytes.49,50 The assay is used for the routine monitoring of patients at high risk for severe CMV disease, including recipients of solid-organ and bone marrow transplants and individuals infected with HIV. The assay can be used to predict and differentiate CMV disease from asymptomatic infection, monitor the efficacy of antiviral therapy and predict drug resistance, and to make decisions regarding the initiation of preemptive therapy.

Solid-Phase Immunoassays

Solid-phase immunoassays also can be used for the detection of viral antigens from clinical specimens.30,48,51 A number of commercial kits are available and include conventional enzyme-linked immunoassays (EIAs) and the more rapid and less sophisticated immunoassays. In conventional EIAs, specific viral antigens, if present in a clinical specimen, will bind to monoclonal antibodies coated onto the surface of a solid phase (e.g., a well of a microtiter plate or a polystyrene bead). Following a wash step, an enzyme-labeled antiviral monoclonal antibody is added. The antibody–antigen–antibody complex is then detected by the addition of a colorless substrate, which becomes colored in the presence of the enzyme. The assays usually take 1–2 hours to complete and the results are read in a spectrophotometer. This type of assay is normally performed in the laboratory and is primarily used for the detection of viral antigens of HBV, rotavirus, adenoviruses, including types 40 and 41, and norovirus.52,53

The more recently developed rapid immunoassays involve self-contained, disposable devices and only a single step or a few simple steps. There are several basic formats, including membrane flow-through devices, lateral-flow immunochromatographic strips, and optical immunoassays. An endogenous enzyme assay also has been commercialized for the direct detection of neuramindase activity of influenza viruses from clinical specimens. The rapid assays require no specialized equipment and require little technical expertise and can be completed in 15–30 minutes. Kits are designed to be used either in the laboratory or at the site where the specimen was collected (e.g., physician's offices, ambulatory clinics, and emergency departments). Rapid assays are available for RSV, influenza virus types A and B, and rotavirus. Although simple and relatively inexpensive to perform, rapid antigen tests are the least accurate of all direct detection tests offered in a diagnostic virology laboratory. A number of false-negative and false-positive results can be generated from these tests depending on how, when, and where the tests are used. This is particularly true for rapid antigen tests for influenza virus, which may vary in sensitivity and specificity depending on the age of the patient, specimen type and adequacy, virus subtype, prevalence of the virus in the community, and the particular test that is selected for testing.51

Molecular Methods

There has been enormous enthusiasm for the medical and commercial potential of molecular technologies, and in the past two decades, there has been an explosion of technological innovations in molecular diagnostics. The development of rapid and sensitive molecular amplification methods has resulted in one of the most dynamic and dramatic revolutions in clinical laboratory medicine, particularly in the diagnosis of viral diseases. A large and growing number of viruses can now be detected using such methods as PCR, nucleic acid sequence-based amplification, branched chain signal amplification, transcription-mediated amplification, hybrid capture signal amplification, and strand displacement amplification. The sensitivity and specificity of these assays exceed that of more conventional methods in the diagnostic virology laboratory and the clinical applications of these techniques seem endless. As such, the emphasis of the clinical virology laboratory is changing considerably.

The 1990s saw a new wave of change that is still going on today with the advent of real-time quantitative PCR,54–58 advancements in microfluidics and microelectronics, and the development of sequencing systems, microarrays, biochips, and biosensors.59–62 Molecular amplification methods are now rapidly displacing the more traditional culture- and antigen-based procedures that have been used for decades and are becoming the new “gold standard” for detecting most viruses of medical importance. These methods can detect viruses for which existing tests are considerably less accurate or for which no tests exist.63 The technologies are being used successfully to detect unculturable, fastidious, or slow-growing viruses and for detecting viruses that are new or otherwise too dangerous to grow. Molecular amplification methods are especially well suited for detecting viruses present in small specimen volumes or that are in low numbers or nonviable within clinical specimens. Multiplex procedures have been developed and commercialized for the simultaneous detection of multiple viruses from a single specimen.64–70 Quantitative molecular amplification assays have become invaluable tools to assess disease progression and prognosis, monitor therapy, predict treatment failure and the emergence of drug resistance, and to facilitate our understanding of the transmission and pathogenesis of certain viruses in chronically infected and immunocompromised hosts. Commercial and user-developed assays are now available for the accurate quantification of viral nucleic acids of HIV-1, CMV, EBV, human herpes virus-6 (HHV-6), HHV-7, HHV-8, BK virus (BKV), HBV, and HCV.71–76 Lastly, molecular genotyping assays that involve using nucleic acid amplification of specific viral genes and direct sequencing of the amplified products have been developed.74,77 These methods are primarily being used to identify mutations that confer resistance to antiviral drugs used for the treatment of HIV-1 and CMV and for recognition of genetic variants of HBV and HCV that may be refractile to antiviral drugs. Use of genotypic assays also can provide valuable information about the evolution and phylogenetic relationships among closely related viruses and the epidemiological and pathogenic behavior of viruses.

The continued development of molecular amplification procedures has led to an explosive increase in the availability of high-quality commercial reagents. Nucleic acid amplification methods are now an integral and necessary component of many diagnostic virology laboratories, and continuous improvements, automation, and simplification of the technology have made these procedures easier to use and more accessible to laboratories with even limited experience. More recent advances have resulted in new generations of rapid molecular amplification assays for detection, quantification, and typing of viruses. This has greatly increased our ability to accurately detect and monitor viral infections. With the continued arrival of more cost-effective and automated nucleic acid isolation and amplification systems, the future holds great promise for the widespread use of such methods in every clinical virology laboratory. Ultimately, the acceptance of these tests will depend on their clinical performance, convenience, and relative expense. The technology will continue to advance and have even a greater impact on the care and management of ill patients with viral infections. However, enthusiasm for the use of molecular-based technologies must be tempered by recognition of the need for performing rigorous quality control in the laboratory and providing appropriate interpretation of results. The significance of the results must be evaluated with respect to the virus identified, the specimen site, and the clinical situation. Also, there must be a greater availability of assays licensed by the U.S. Food and Drug Administration (FDA) as there are only a few FDA-cleared commercial molecular test kits in the market for laboratories to use. As such, many laboratories have been forced to develop their own molecular assays, thereby limiting much of the molecular testing to laboratories at academic institutions or large reference laboratories with the expertise, personnel, and resources to enable these technologies.

Cytology/Histology

Direct cytological or histological examinations of stained clinical material are some of the fastest and oldest methods of detecting viruses. While relatively simple and cost-effective, the tests are insensitive compared with direct antigen or nucleic acid detection methods. The specificity can also be low; Tzanck smears, for example, are limited by their inability to distinguish HSV from VZV infections. Cytologic examination of exfoliated cells has been applied to specific viruses such as CMV, HSV, VZV, adenovirus, the polyomaviruses BK and JC, measles virus, rabies virus, and human papillomaviruses (HPV). Histological examination of impression smears, frozen sections, or formaldehyde-fixed and paraffin-embedded tissue has been used for various viruses, including CMV, EBV, HSV, BKV, HBV, HPV, parvovirus B19, and adenovirus, and may provide useful information regarding tissue inflammation and damage as the result of viral infection. The sensitivity of histological staining can be increased somewhat by using immunohistochemical or in situ hybridization techniques. Overall, these tests are used sparingly in most laboratories.

Electron Microscopy

Electron microscopy can be a useful tool for the rapid identification of viral particles based on characteristic size and morphology.78,79 It offers the main advantage of speed when doing negative staining of liquid samples and can detect fastidious or uncultivable viruses. The method has been mostly applied to the examination of stools for viral agents of gastroenteritis and has been used successfully as an adjunct to other methods for detecting unidentified viruses suspected of causing disease. The major limitations include the high cost of the instrument, the requirement for specialized facilities and expertise, and moderate to low sensitivity and specificity. This procedure has largely been replaced by alternative methods for viral diagnosis and is seldom available in diagnostic laboratories in the United States.

Serology

A number of sensitive and specific tests are available for the detection of antibodies to a variety of viruses.80 Enzyme immunoassays, immunofluorescence, or passive latex agglutination tests are commonly used by most laboratories to screen for the presence of viral-specific antibodies in a clinical specimen. Immunoblot techniques are available for HIV-1 and -2, HCV, and human T-cell leukemia virus-I and -II (HTLV-I and -II), and are primarily used as confirmatory or supplemental tests to verify the results of positive screening tests. Serological testing can be useful for the diagnosis of recent or chronic viral infections and to determine the immune status of an individual or group. Antibody detection remains at the forefront of diagnosis of infections with HIV-1 and -281 and the hepatitis viruses A–E,82 as well as EBV, the arboviruses, measles, mumps and rubella viruses, parvovirus B19, and HTLV-I and -II. Defining an individual's immunity to a given virus can be beneficial for (1) prenatal and pretransplanatation screening, (2) testing blood and blood products for donation, (3) postexposure monitoring, (4) preemployment screening in a patient care setting, and (5) verifying an immune response following administration of vaccines. Detection of virus-specific IgM in a single serum sample can be diagnostic of primary viral infection and has been used successfully for many viruses. Viruses for which IgM testing can be useful include CMV, EBV, VZV, HHV-6 and -7, measles, mumps and rubella viruses, hepatitis A virus (HAV), HBV, parvovirus B19, and arthropod- and rodent-borne viruses. Seroconversion from a negative to a positive IgG antibody response between acute and convalescent sera collected 2–3 weeks apart can also be used to diagnose a primary infection, but such testing is no longer routinely performed in most hospital diagnostic laboratories since it is retrospective and has a limited impact on the care and management of patients. Detection of virus-specific IgG in a single serum specimen indicates exposure to a virus at some time in the past or a response to vaccination, while finding no detectable antibodies may exclude viral infection. Results of serological tests must be interpreted with caution, as measurements and interpretations of antibody responses to viral infections can be complicated by numerous factors.80 For most viral infections in the acute phase of illness, rapid antigen and/or nucleic acid detection methods or viral isolation are also available and may yield results in a more sensitive and timely manner.

Rapid and simple tests for the detection of HIV antibodies have been developed and licensed by the FDA.83–85 These assays involve no special equipment, require little technical expertise, and are performed using self-contained, disposable devices. Most of the assays use serum or plasma for testing, while whole blood and oral fluids have also been incorporated as acceptable specimens for some assays. The assays have been designed to detect HIV-1 only or both HIV-1 and -2, and can be performed at the point of care or in the laboratory. The sensitivity and specificity of the rapid assays are comparable to laboratory-based screening tests, and like laboratory-based assays, confirmation by Western blot or immunofluorescence is required for specimens that are positive for HIV-specific antibody by rapid testing. Rapid HIV tests have been used widely in developing countries as tools for screening and confirmation of an HIV antibody response. They are the preferred test in this setting since resources and facilities may not be available to perform the more technically demanding laboratory-based immunoassays and Western blots. In the United States, these tests are being advocated for use in emergency departments, hospital clinics, sexually transmitted disease (STD) clinics, family planning clinic, and outreach programs. The intended uses of rapid HIV tests include providing greater access to testing and counseling and same-visit results, screening pregnant women with unknown HIV serostatus at the time of delivery, and assessing the risk of HIV transmission following exposure.

More so than any other time in the history of diagnostic virology, a number of rapid and accurate laboratory tests are now available to health care providers faced with children who are acutely or chronically ill with viral diseases. The appropriate selection, use, and interpretation of these methods in combination with clinical assessment of the patient can greatly improve care and have a positive impact on the management of these ill patients.