Immunoassays (Molecular Biology)

Immunoassays are a collection of techniques for measuring biological or synthetic materials, using antibodies specific for those materials. Immunoassays are also applied in reverse, in which a purified antigen or hapten is used to quantify an uncharacterized antibody preparation. The common element in all immunoassays is the interaction of an antibody with a ligand. The assay methods differ in the accessory chemical reactions that take place prior to, or as a consequence of, ligand binding, and in the instrumentation used to detect the binding event or subsidiary reaction. Radioimmunoassays and enzyme-linked immunosorbent assays (ELISA) are especially widely used types of immunoassay. Others are complement fixation, agglutination, Farr Assay, and Ouchterlony Double Diffusion. They are listed as separate entries and not discussed here. Below, we sketch the principles underlying the most common immunoassay methods relevant to molecular biology.

1. Fluorescence Quench

The emission of a photon as a molecule returns to its electronic ground state from an excited state produces fluorescence. "Quenching of fluorescence" refers to chemical interactions that allow an excited state to dissipate its energy through a nonradiative mechanism (see Fluorescence Quenching). Common mechanisms that can cause quenching include collisions with solvent, protonation or deprotonation reactions, resonance energy transfer, and collisions or static contact with quenchers. Quenchers encompass a huge variety of substances, from molecular oxygen to heme proteins, and the binding of a quencher to an antibody can easily be measured. Quenchers most relevant to immunoassays are aromatic organic molecules, which are frequently used as haptens. These molecules are thought to cause nonradiative deexcitation by transiently accepting electrons from the excited state of a fluorophore.

Tryptophan is the only strongly fluorescent naturally occurring amino acid, and antibody sequences usually show an abundance of exposed Trp residues in the antigen-combining site (1). Consequently, binding of a quenching antigen or hapten to an antibody nearly always causes decrease of Trp fluorescence from the antibody. Furthermore, the fluorescence decrease is generally linear with the number of combining sites filled; hence it is a direct measure of the degree of saturation of the antibody (2). Bound and free Ab as a function of added ligand can be calculated and displayed as a Scatchard Plot or used for direct fitting to a binding equation to determine the antibody valency and the association equilibrium constant for binding the ligand. Fluorescence quench on antigen binding is a very desirable technique for immunoassay, because it allows direct measurement of antibodies and antigens interacting in solution and can be performed without chemical labeling of the molecules under study with reporter groups.

2. Fluorescence Anisotropy or Fluorescence Polarization

This method relies on the change in molecular tumbling rate that occurs when an antibody-antigen complex forms. Fluorescence anisotropy is most appropriate for antibody-antigen pairs that are of different molecular size (3), and may be applicable even if no overt change in fluorescence signal results from complex formation.

Plane-polarized light absorbed by a fluorophore will (to a first approximation) be emitted with the same plane of polarization, unless the fluorophore molecule reorients during the lifetime of its excited state. Since the tumbling rate is a function of molecular size, polarization of emitted light is very sensitive to complex formation. A small molecule will tumble rapidly, hence the orientation of an excited state fluorophore will randomize, and fluorescence emission will be isotropic. In contrast, a large molecule will tumble slowly; hence emission from a fluorophore will remain polarized. A small molecule that binds to a large molecule will take on the tumbling characteristics of the large molecule. For example, anisotropic emission from a fluorescent tag on a small antigen will sharply increase on complexation with a large antibody. To use this principle for a quantitative immunoassay, fluorescence anisotropy is measured separately for the free antigen and fully complexed antigen. Between these two limiting values, anisotropy will be linear with the extent of antigen complexation; hence the degree of complex formation in a test sample may be inferred from the observed anisotropy.

Fluorescence anisotropy is measured by exciting a fluorophore with polarized light and observing fluorescence intensity through a polarizing filter oriented parallel (/y;) or perpendicular (/,_;) to the excitation plane. The relative intensities from the parallel and perpendicular emissions are used to calculate anisotropy or polarization from the equations:

Although polarization and anisotropy are very closely related, equations relating observed intensities to ligand binding and other phenomena are generally simpler when expressed in terms of anisotropy; hence this is the preferred parameter. Fluorophores used for anisotropy measurements of proteins, such as fluorescein and dansyl, typically have excited-state lifetimes of 5 ns. This lifetime is on the same order as the tumbling times of protein antigens. The small, globular protein lysozyme, for example, also has a rotational correlation time of 5 ns (4); hence a fluorescein label on lysozyme would show considerable depolarization of fluorescence. Intact IgG has a rotational correlation time of 220 ns; hence its complex with the labeled lysozyme would show strong fluorescence anisotropy.

One complication to the fluorescence anisotropy technique is the occurrence of local motions affecting the fluorophore, often referred to as "segmental flexibility." The timescale for these local motions can be sufficiently fast to reorient the fluorophore and depolarize emission, even if the fluorophore is attached to a large, slowly tumbling macromolecule (5).

3. Phosphorescence or Time-Resolved Fluorescence

Phosphorescence is a fluorescence phenomenon in which, for quantum-mechanical reasons, the excited state of a fluorophore cannot easily return to the ground state by emission of a photon. Emission does occur, but on a much longer timescale than is normal for fluorescence. Because of this long lifetime, the presence of even small amounts of quenching agents such as O 2 is sufficient to suppress observation of any signal from naturally phosphorescent biological compounds. Phosphorescence is measured by exciting a sample with a pulsed light source, waiting an interval for short-lived fluorescence states to decay, then integrating photon counts during the remaining dark period before the next pulse. This ability to resolve short-lived from long-lived fluorescence is the chief advantage of a phosphorescent probe, as background phosphorescence of biological samples is almost nil. By contrast, the steady-state fluorescence background is often quite high, and the signal-to-background ratio in short-lived fluorescence-based assays is a more common limitation of sensitivity of an assay than the absolute magnitude of the signal.

Europium is the most common phosphorescent probe in time-resolved fluorescence immunoassays (6). This metal ion is attached to antibodies or antigens through bifunctional linker molecules containing a chelate moiety at one end and a group reactive with protein residues at the other. In a typical assay, a trapping antibody is immobilized on a solid support, antigen in a test sample is allowed to bind, and then the Eu -labelled antibody is allowed to bind the trapped antigen. After washing away unbound reagents, Eu is extracted from the antibody in low pH buffer containing phosphorescence enhancing agents (7). These agents act both by sequestering the Eu ion from quenchers and transferring absorbed light to the Eu ion. The fluorescence emission is integrated repetitively over several hundred microseconds, using a stroboscopic fluorimeter. Comparison of the response from the test sample to a standard curve constructed with known concentrations of analyte allows determination of the concentration of analyte in the test sample.

4. Chemiluminescence

Chemiluminescence is a phenomenon related to fluorescence, in which an electronic excited state of a molecule is reached through a chemical reaction. Emission of a photon, which is optically detected, returns the molecule to the ground state. A chemiluminescent immunoassay involves an antibody or antigen that has been covalently tagged with a molecule capable of participating in a chemiluminescent reaction. The tag remains inert through immunologic steps such as antigen or secondary antibody binding. Subsequent mixing with a trigger solution initiates the luminescent reaction.

A chemiluminescent immunoassay to detect an antigen typically begins with immobilization of a capture antibody on a solid support. The test sample is added, followed by the tagged detection antibody. Luminol was the first luminogenic tag to be used for an immunoassay (8), but derivatized acridinium esters are now more common (9, 10). Unbound reagents are washed away, and luminescence is determined in a luminometer. This device combines two automated functions (11). First, trigger solutions (alkaline H 2O2 in the case of acridinium labels) are added to the sample to initiate the luminescence reaction. Mixing is usually complete in less than 1 s. Next, a detector, usually with photon-counting electronics, measures light emission from the sample over a defined time interval, typically 10 s. As with other immunoassays, the response of a test sample is compared to the responses from a set of standards to determine the analyte concentration in the test sample.

Because photon emission from luminescent tags can be induced to occur quantitatively over the span of a minute, as compared to radioactive decay of isotopic isotopes, which continues over months in 125 the case of I, chemiluminescent assays are inherently very sensitive. ELISA assays that use a luminogenic substrate, which are sometimes referred to as "chemiluminescent immunoassays," are perhaps the most sensitive of all assays. Combination of an enzyme tag on the antibody and a luminogenic enzymatic reaction yields a degree of amplification that can detect as little as 1000 molecules of analyte (12).