Principle

Formation of a macromolecular complex is a fundamental prerequisite for nephelometric protein quantitation. The procedure is based on the reaction between the protein being assayed and a specific antiserum. Protein in a patient’s specimen reacts with specific nephelometric antiserum to human proteins and forms insoluble complexes. When light is passed through such a suspension, the resulting complexes of insoluble precipitants scatter incident light in solutions. The scattered light can be detected with a photodiode. The amount of scattered light is proportional to the number of insoluble complexes and can be quantitated by comparing the unknown patient values with standards of known protein concentration.

The relationship between the quantity of antigen and measuring signal at a constant antibody concentration is expressed by the Heidelberger curve. If antibodies are present to excess, a proportional relationship exists between the antigen and resulting signal. If the antigen overwhelms the quantity of antibody, the measured signal drops.

By optimizing the reaction conditions, the typical antigen-antibody reactions as characterized by the Heidelberger curve are effectively shifted in the direction of high concentration. This ensures that these high concentrations will be measured on the ascending portion of the curve. At concentrations higher than the reference curve, the instrument will transmit an out of range warning.

Physical Basis

Nephelometry is based on the principle that light is scattered by a homogeneous particulate solution at a variety of angles. Three types of scatter can occur: (1) scatter around the particles; (2) forward scatter caused by out of phase backscatter; and (3) forward scatter exceeding backscatter.

Optical System

In the nephelometric method, an infrared high-performance, light-emitting diode (LED) is used as the light source. Because an entire solid angle is measured after convergence of this light through a lens system, an intense measuring signal is available when the primary beam is blocked off. In connection with the lens system, this produces a light beam of high colinearity. The wavelength is 840 nm. Light scattered in the forward direction in a solid angle to the primary beam ranges between 13 and 24 feet and is measured by a silicon photodiode with an integrated amplifier. The electrical signals generated are digitized, compared with reference curves, and converted to protein concentrations.

Measuring Methods

A fixed-time method is used routinely for precipitation reactions. Ten seconds after all reaction components have been mixed, a cuvette reading (initial blank measurement) is taken. A second measurement is taken 6 minutes later and, after subtraction of the original 1-second blanking value, a final answer is calculated against the multiple-point or single-point calibration in the computerized program memory for the assay.

Advantages and Disadvantages

Nephelometry represents an automated system that is rapid, reproducible, relatively simple to operate, and common in higher volume laboratories. It has many applications in the immunology laboratory. Currently, instruments using a rate method and fixed-time approach are commercially available with tests for immunoglobulin G (IgG), IgA, IgM, C3, C4, properdin, C-reactive protein (CRP), rheumatoid factor, ceruloplasmin, α₁-antitrypsin, apolipoproteins, and haptoglobins.

The disadvantages of nephelometry include high initial equipment cost and interfering substances such as microbial contamination, which may cause protein denaturation and erroneous test results. Intrinsic specimen turbidity or lipemia may exceed the preset limits. In these cases, a clearing agent may be needed before an accurate assay can be performed. In addition, low-molecular-weight immunoglobulins, monoclonal immunoglobulins, and antibovine antibodies also may produce spurious results in nephelometry.

Clinical Application: Cryoglobulins

Cryoglobulin analysis is frequently requested when patient symptoms such as pain, cyanosis, Raynaud’s phenomenon, and skin ulceration on exposure to cold temperatures are present. Cryoglobulins are proteins that precipitate or gel when cooled to 0° C (32° F) and dissolve when heated. In most cases, monoclonal cryoglobulins are IgM or IgG. Occasionally, the macroglobulin is both cryoprecipitable and capable of cold-induced anti-i–mediated agglutination of red blood cells.

Cryoglobulins with a detected monoclonal protein component normally prompt a clinical investigation to determine whether an underlying disease exists. Cryoglobulins are classified as follows:

To test for the presence of cryoglobulins, blood is collected, placed in warm water, and centrifuged at room temperature. The serum is then put into a graduated centrifuge tube and placed in a 4° C (39° F) environment for 7 days. If a gel or precipitate is observed, the tube is centrifuged and the precipitate is washed at 4° C (39° F), redissolved at 37° C (98.6° F), and evaluated by double diffusion and immunoelectrophoresis for the content of the cryoglobulin. Newer methods use nephelometry with cold treatment for analysis.

Principles of Cell Cytometry

Flow cell cytometry, developed in the 1960s, combines fluid dynamics, optics, laser science, high-speed computers, and fluorochrome-conjugated monoclonal antibodies (MAbs) that rapidly classify groups of cells in heterogeneous mixtures. The principle of flow cytometry is based on cells being stained in suspension with an appropriate fluorochrome—an immunologic reagent, a dye that stains a specific component, or some other marker with specific reactivity. Fluorescent dyes used in flow cytometry must bind or react specifically with the cellular component of interest (e.g., reticulocytes, peroxidase enzyme, DNA content). Fluorescent dyes include acridine orange and thioflavin T. Pygon is preferred for fluorescein isothiocyanate (FITC) labeling. Krypton is often used as a second laser in dual-analysis systems and serves as a better light source for compounds labeled by tetramethyl-rhodamine isothiocyanate and tetramethylcyclopropyl-rhodamine isothiocyanate.

A suspension of stained cells is pressurized using gas and transported through plastic tubing to a flow chamber within the instrument (Fig. 13-3). In the flow chamber, the specimen is injected through a needle into a stream of physiologic saline called the sheath. The sheath and specimen both exit the flow chamber through a 75-µm orifice. This laminar flow design confines the cells to the center of the saline sheath, with the cells moving in single file.

The stained cells then pass through the laser beam. The laser activates the dye and the cell fluoresces. Although the fluorescence is emitted throughout a 360-degree circle, it is usually collected by optical sensors located 90 degrees relative to the laser beam. The fluorescence information is then transmitted to a computer, which controls all decisions regarding data collection, analysis, and cell sorting.

Flow cytometry performs fluorescence analysis on single cells. The major applications of this technology are as follows:

Immunophenotyping

Monoclonal antibodies, identified by a cluster designation (CD), are used in most flow cytometry immunophenotyping (Table 13-1). Cell surface molecules recognized by monoclonal antibodies are called antigens because antibodies can be produced against them or are called markers because they identify and discriminate between (mark) different cell populations. Markers can be grouped into several categories. Some are specific for cells of a particular lineage (e.g., CD4+ lymphocytes) or maturational pathway (e.g., CD34+ progenitor stem cells); the expression of others can vary, according to the state of activation or differentiation of the same cells.

In flow cytometry, cells can be sorted from the main cellular population into subpopulations for further analysis (Fig. 13-4). Any fresh specimen that can be placed into a single-cell suspension is a valid candidate for immunophenotyping (e.g., T cells, B cells, CD34+ stem cells; detection of minimal residual disease in leukemia). Sorting is accomplished using stored computer information.

When the laser strikes a stained cell, the dye creates distinctive colored light that the cytometer recognizes. This fluorescent intensity is recorded and analyzed by the computer and cells are sorted according to a preprogrammed selection. If the particular cell in the laser beam is of interest, the computer waits the appropriate time for the cell to reach the droplet break-off point within the charging collar. At that point, the computer signals the charging collar to administer an electrostatically positive or negative charge to the stream containing the target cell. A droplet containing this cell is then removed from the main stream before the charge has time to redistribute.

This action produces the cell of interest within a liquid drop that has an electrostatic charge on its surface (only the droplet is charged). The droplet falls between a set of deflection plates, which creates an electrical field. The charged droplets are deflected to the left or right, depending on their polarity, and collected for further analysis.

Multicolor Immunofluorescence

Current fluorescent methods (e.g., BD FACSCanto II flow cytometer; BD, Franklin Lakes, NJ) can perform up to eight-color analysis. The BD LSRII flow cytometer, with up to four lasers, can measure up to 16 colors. It can use four MAbs, each directly conjugated to a distinct fluorochrome, per tube of patient cell suspension. The four most common fluorochromes are FITC, phycoerythrin (PE), peridinin chlorophyll protein (PerCP), and allophycocyanin (APC). The first three fluorochromes are excited by the 488-nm line of an argon laser; the fourth fluorochrome is excited by the 633-nm line of a helium-neon or diode laser.

Eight-color immunofluorescence offers the advantages of greater sensitivity and specificity, with increased ability to identify and subclassify individual cells. Improvements in methods and probes may lead to fluorescence in situ hybridization (FISH) in suspension as a routine protocol (see Chapter 12) and enable flow cytometry to operate on a molecular level simultaneously to identify chromosomal abnormalities.

A system that uses a flow cytometer, specific data analysis software, and fluorescent latex particles, the Luminex 100 Total System, has been developed by Luminex Technology (Austin, Texas). This system combines advances in computing and optics with a new concept in color coding to create a simple, cost-effective analysis system (Fig. 13-5). Latex beads are coupled to various amounts of two different fluorescent dyes, which are analyzed by the flow cytometer and software to allow the distinct separation of up to 64 slightly different colored bead sets. The color-coded microspheres identify each unique reaction. Hundreds of microsphere sets can be identified at once in a single sample. Optical technology recognizes each microsphere and provides a precise, quantitative measure simply and in real time.

Currently, up to 64 microsphere sets are recognized. The current FlowMetrix system is compatible with the BD FACS Vantage SE System and BD FACSCalibur, the most widely used flow cytometers for cellular analysis. Because Luminex technology requires fewer steps to assess multiple parameters, with a high level of sensitivity and accuracy, it is significantly more cost-effective than current methods of analysis. Some immunologic applications already demonstrated with FlowMetrix are human immunodeficiency virus (HIV) and hepatitis B seroconversion, multicytokine measurement, multiplexed allergy testing, DNA-based tissue typing, herpes simplex viral load, IgG, IgA, and IgM assays, IgG subclassification, autoimmunity panel, epitope mapping, human chorionic gonadotropin (hCG) and α-fetoprotein, HIV viral load, and the TORCHS (toxoplasmosis, other [viruses], rubella, cytomegalovirus, herpesviruses, syphilis) panel.

Sample Preparation

Specimens that can be used for flow cell analysis include whole blood, bone marrow, and aspirates of body fluids. Whole blood, collected in ethylenediaminetetraacetic acid (EDTA), is the preferred anticoagulant if specimens are processed within 30 hours of collection. Heparin is an alternative anticoagulant for whole blood and bone marrow and can provide stability of specimens more than 24 hours old.

Blood specimens should be stored at room temperature (20° C to 25° C [68° F to 77° F]) before processing. Specimens need to be well mixed prior to delivery into staining tubes. Unsuitable specimens included hemolyzed or clotted samples. For the efficient analysis of white blood cells, whole blood, bone marrow, or aspirates should have the bulk of red blood cells removed prior to analysis. Tissue specimens (e.g., lymph nodes) should be collected and transported in a tissue culture medium at room temperature or at 4° C (39° F) if analysis is delayed. Such a specimen requires disaggregation by enzymatic or mechanical methods to form a single-cell suspension. After proper specimen processing, antibodies are added to the cellular preparation and analyzed. MAbs, tagged with different fluorescent tags, are used for analysis.

A six-color flow cytometry diagnostic application uses the BD FACSCanto II flow cytometer and BD Multitest six-color TBNK with BD Trucount tubes to determine the absolute counts of mature T, B, and natural killer (NK) lymphocytes (Fig. 13-6), as well as CD4+ and CD8+ T cell subsets in human peripheral blood, in a single tube.

Fluorescent Polarization Immunoassay

The fluorescent polarization immunoassay IMX System (Biostad-Abbott, Quebec) is an automated analyzer designed to perform microparticle enzyme immunoassay and fluorescence polarization immunoassay using ion capture technology. This unique combination allows both high- and low-molecular-weight analytes to be measured. This expands the range of available assays to include tests for endocrine function, fertility, cancer, hepatitis, transplantation, rubella, and congenital disease.

Animated Flow Cytometry Theory—Theory #1

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Animated Flow Cytometry Theory—Theory #5

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