Principles of Virology. Jane Flint

may also be detected in infected animals by immunohistochemistry. In this procedure, tissues are embedded in a solid medium such as paraffin, and thin slices are produced using a microtome. Viral antigens can be detected within the cells in the sections by direct and indirect immunofluorescence assays.

Enzyme immunoassay. Detection of viral antigens or antiviral antibodies can be accomplished by solid-phase methods, in which an antiviral antibody or protein is adsorbed to a plastic surface (Fig. 2.13A). To detect antibodies to viruses, viral protein is first linked to the plastic support, and then the specimen is added (Fig. 2.13B). Like other detection methods, enzyme immunoassays are used in both experimental and diagnostic virology. In the clinical laboratory, enzyme immunoassays are used to detect a variety of viruses, including rotavirus, herpes simplex virus, and human immunodeficiency viruses. A modification of the enzyme immunoassay is the lateral fow immunochromatographic assay, which has been used in rapid antigen detection test kits (Fig. 2.14). The lateral fow immunochromatographic assay does not require instrumentation and can be read in 5 to 20 min in a physician’s office or in the field. Commercial rapid antigen detection assays are currently available for influenza virus, respiratory syncytial virus, and rotavirus.

Figure 2.13 Detection of viral antigen or antibodies against viruses by enzyme-linked immunosorbent assay (ELISA). (A) To detect viral proteins in serum or clinical samples, antibodies specific for the virus are immobilized on a solid support such as a plastic well. The sample is placed in the well, and viral proteins are “captured” by the immobilized antibody. After washing to remove unbound proteins, a second antibody against the virus is added, which is linked to an indicator. The second antibody will bind if viral antigen has been captured by the first antibody. Unbound second antibody is removed by another washing, and when the indicator is an enzyme, a chromogenic molecule that is converted by the enzyme to an easily detectable product is then added. The enzyme amplifies the signal because a single catalytic enzyme molecule can generate many product molecules. Another wash is done to remove unbound second antibody. If viral antigen has been captured by the first antibody, the second antibody will bind and the complex will be detected by the indicator. (B) To detect antibodies to a virus in a sample, viral antigen is immobilized on a solid support such as a plastic well. The test sample is placed in the well, and antiviral IgG antibodies present in the sample will bind the immobilized antigen. After washing to remove unbound components in the sample, a second antibody, directed against a general epitope on the first antibody, is added. Unbound second antibody is removed by another wash. If antibodies against the virus are present in the specimen, the second antibody will bind to them and the complex will be detected via the indicator attached to the second antibody, as described in (A).

       Fluorescent Proteins

The discovery of green fluorescent protein revolutionized the study of the cell biology of virus infection. This protein, isolated from the jellyfish Aequorea victoria, is a convenient reporter for monitoring gene expression, because it is directly visible in living cells without the need for fixation, substrates, or coenzymes. Mutagenesis of the gene encoding this protein has led to the development of new fluorescent probes ranging in color from blue to yellow (Fig. 2.15A). Additional fluorescent proteins emitting in the red, deep red, cyan, green, yellow, and orange spectral regions have been isolated from other marine species. Codon optimization for maximum translation in specific cell types and improved stability and brightness are other modifications that have broadened the utility of these proteins.

       Fluorescence Microscopy

      Fluorescence microscopy allows virologists to study all steps of virus reproduction, including cell surface attachment, cell entry, trafficking, replication, assembly, and egress. Single virus particle tracking can be achieved by inserting the coding sequence for a fluorescent protein into the viral genome, often fused to the coding region of a viral protein. The fusion protein is incorporated into the viral particle, which is visible in cells by fluorescence microscopy (Fig. 2.15B). An alternative approach is to attach small-molecule fluorophores to viral capsid proteins. Light microscopy has a resolution in the range of 200 to 500 nm, whereas most viruses are between 20 and 400 nm in size and are therefore below the difraction limit. However, when the virus particle emits a high fluorescent signal in a low background, it is possible to use a computational point tracking algorithm to locate the particle with greater precision than the diffraction limit of the light microscope. This technique allows single particle tracking with accuracy in the range of low tens of nanometers.

      Recent improvements in microscopy technology and computational image manipulation have led to unprecedented levels of resolution and contrast and an ability to reconstruct three-dimensional structures from captured images. The first advance was confocal microscopy, which utilizes a scanning point of light instead of full-sample illumination. In a conventional light microscope, light can penetrate the specimen only to a fixed depth. In a confocal microscope, a small beam of light is focused to multiple narrow depths. By capturing multiple two-dimensional images at different depths, it is possible to reconstruct high-resolution three-dimensional structures, a process known as optical sectioning.

Figure 2.14 Lateral flow immunochromatographic assay. A slide or “dipstick” covered with a membrane is used to assay for the presence of viral antigens. The clinical specimen is placed on an absorbent pad at one end and is drawn across the slide by capillary action. Antigens in the sample react with a virus-specific antibody, which is linked to an indicator, in this example, colloidal gold. The antigen-antibody complexes move across the membrane until they are captured by a second virus-specific antibody in a test line. If viral antigen is present in the sample, an indicator line becomes visible in the test line. Accumulation of the indicator-containing antibody at the control line provides validation that the assay is functioning.

      Superresolution microscopy combines the advantages of fluorescent imaging (multicolor labeling and live-cell imaging) while breaking the resolution limit of light microscopy. Different formats include single molecule localization microscopy, in which only a subset of fluorophores are turned on during each imaging cycle, thus allowing position determination with nanometer accuracy. Fluorophore positions from a series of images are then used to reconstruct the final image. Structured illumination microscopy utilizes standing waves formed by interference in laser illumination to create an excitation field that allows optical sectioning at very high resolution. These approaches can achieve resolution below 1 nm, well below the limit of light microscopy. This resolution is achieved by combining sequential acquisition of images with random switching of fluorophores on and off. From several hundred to thousands of images are collected and processed to generate a superresolution data set that can resolve cellular ultrastructure.

      These superresolution microscopy methods are well suited for providing high-resolution images of static sections. Because these methods acquire images slowly, are phototoxic, and require computationally intensive image processing, their use for time-lapse imaging of live cells is impractical.

      Fluorescence resonance energy transfer (FRET) microscopy can be used to examine protein-protein and protein-DNA or RNA interactions and conformational changes in these molecules. FRET solves the problem encountered in conventional