With the development of science and technology, a growing numbers of biological products are applied clinically in human bodies. Meantime, there is a concern for regulatory agencies and manufacturers about the immunogenicity of biological products because immune response can lead to different kinds of effects for the effectiveness of products. And immunogenicity of biological products can occur pre-clinically and clinically when products induce immune responses in animals or humans receiving the products. A number of factors induce the immune system responding to the presence of a product, such as protein structure, glycosylation, contaminants, formulation and degradation products. (1)Various kinds of clinical results of immunogenicity can lead to various kinds of effects which can range from no effects to very serious adverse effects even the death. But it is very rare to happen. Whilst there are many methods to detect immunogenicity, they depend on detecting the body fluid instead of the cellular response of the immune system. And there are different assays such as immunoassays or bioassays are widely used in the detection of immunogenicity. The design of assays are critical to assess the immunogenicity. These computational and laboratory-based methods for the prediction of immunogenicity can reduce potential immunogenicity of biological products which will induce less immunogenicity of biological products and increase the products safety in future. (1)
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Biological products (also called biologics) are made from a variety of natural sources such as human, animal or microorganism. Like drugs, some biologics are used to treat diseases and medical conditions and prevent or diagnose diseases. Biologics can be made up of sugars, proteins, or nucleic acids or complex combinations of these substances, or livings such as cells and tissues. Examples of biological products include vaccines, blood and blood products, allergenic extracts, human cells and tissues used for transplantation, gene therapies, cellular therapies and tests to screen potential blood donors for infectious agents such as HIV. (2)
Vaccines involves viral vaccines and bacterial vaccines. Viral vaccines have developed recently to treat and prevent many kinds of diseases. These vaccines comprise engineered for RNA viruses, rotavirus, INFLUENZA A and B, human papilloma virus (HPV), and varicella-zoster virus (shingles). Researchers develop vaccines for human immunodeficiency virus (HIV) and hepatitis C through the use of biological products in the future. (3)
Bacterial vaccines contain 3 basic types: killed whole organisms or bacterins, including those for pertussis (whooping cough) and those for use in the veterinary or aquaculture fields; single-antigen vaccines extracted from bacteria or prepared by genetic engineering; and toxoids. Many bacteria, such as those that cause tetanus, pertussis or diphtheria, release toxins that cause cellular damage. These toxins have been purified and inactivated, usually with chemicals to produce toxoids. When injected, toxoids induce the formation of antibodies against the original toxin. Some other bacterial vaccines produced include typhus, typhoid, cholera, haemophilus, influenzae type B, pneumococcus, meningococcus and bacille Calmette-Guérin (BCG, used for the prevention of tuberculosis). Vaccines for veterinary use include those for household pets, farm animals and other cultivated species, such as mink or fish. The vaccination of fish is a new approach necessitated by the crowding of fingerlings in AQUACULTURE operations. Administered by injection, immersion or spray, such vaccines are remarkably effective. (3)
Blood Fractions and Serums was started in the 1930s and stimulated by WWII, which was collected and freeze-dried. When combined with water, this product was used in the treatment of blood loss resulting from wound. Methods of separating blood plasma (noncellular fluid) into its constituent proteins were developed in the US. These permitted the Canadian Red Cross Blood Transfusion Service to expand the applications for donated blood. Products prepared from donated blood include red cells, white cells, platelets, and plasma which is fractionated to albumin, immune serum globulins (including specialized products such as tetanus, Rh and rabies immunoglobulins), and coagulation factor concentrates for the treatment of hemophilias A and B. (3)
Methods to alter immunogenicity
Technology to detect immunogenicity of biological products
This picture shows briefly how to detect the immunogenicity of biological products. According to this process, clearly the immunoassays play a very important role in the detection. And immunoassays are mainly used to detect the binding antibody. The antigen -antibody interaction is a biomolecular association similar to an enzyme-substrate interaction, with an important distinction: it does not lead to an irreversible chemical alteration in either the antibody or the antigen. The association between an anti-body and an antigen involves various noncovalent interactions between the antigenic determinant, or epitope, of the antigen and the variable-region (VH/VL) domain of the antibody molecule, particularly the hypervariable regions,or complementarity-determining regions (CDRs).The exquisite specificity of antigen-antibody interactions has led to the development of a variety ofimmunologic assays, which can be used to detect the presence of either antibody or antigen .Immunoassays have played vital roles in diagnosing diseases, monitoring the level of the humoral immune response, and identifying molecules of biological or medical interest. These assays differ in their speed and sensitivity; some are strictly qualitative, others are quantitative.
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The noncovalent interactions that form the basis of antigen-antibody (Ag-Ab) binding include hydrogen bonds, ionic bonds,hydrophobic interactions, and van der Waals interac-tions. Because these interactions are individually weak, a large number of such interactions are required to form a strong Ag-Ab interaction. Furthermore ,each of these noncovalent interactions operates over a very short distance, generally about 1X10-7mm;consequently,a strong Ag-Ab interaction depends on a very close fit between the antigen and antibody. Such fits require a high degree of complementarity between antigen and antibody, a requirement that underlies the exquisite specificity that characterizes antigen-antibody interactions. The combined strength of the noncovalent interactions between a single antigen-binding site on an antibody and a single epitope is the affinity of the antibody for that epitope. Low-affinity antibodies bind antigen weakly and tend to dissociate readily, whereas high-affinity antibodies bind antigen more tightly and remain bound longer.
Enzyme-linked immunosorbent assay, commonly known as ELISA (or EIA), is similar in principle to RIA but depends on an enzyme rather than a radioactive label. An enzyme conjugated with an antibody reacts with a colorless substrate to generate a colored reaction product. Such a substrate is called a chromogenic substrate. A number of enzymes have been employed for ELISA, including alkaline phosphatase, horseradish peroxidase, and galactosidase. These assays approach the sensitivity of RIAs and have the advantage of being safer and less costly. There Are Numerous Variants of ELISA. A number of variations of ELISA have been developed, allowing qualitative detection or quantitative measurement of either antigen or antibody. Each type of ELISA can be used qualitatively to detect the presence of antibody or antigen. Alternatively, a standard curve based on known concentrations of antibody or antigen is prepared, from which the unknown concentration of a sample can be determined.
Antibody can be detected or quantitatively determined with an indirect ELISA (Figure 6-10a). Serum or some other sample containing primary antibody (Ab1) is added to an antigen- coated microtiter well and allowed to react with the antigen attached to the well. After any free Ab1 is washed away, the presence of antibody bound to the antigen is detected by adding an enzyme-conjugated secondary anti-isotype antibody (Ab2), which binds to the primary antibody. Any free Ab2 then is washed away, and a substrate for the enzyme is added. The amount of colored reaction product that forms is measured by specialized spectrophotometric plate readers, which can measure the absorbance of all of the wells of a 96-well plate in seconds.
Indirect ELISA is the method of choice to detect the presence of serum antibodies against human immunodeficiency virus (HIV), the causative agent of AIDS. In this assay, recombinant envelope and core proteins of HIV are adsorbed as solid-phase antigens to microtiter wells. Individuals infected with HIV will produce serum antibodies to epitopes on
these viral proteins. Generally, serum antibodies to HIV can be detected by indirect ELISA within 6 weeks of infection.
Antigen can be detected or measured by a sandwich ELISA (Figure 6-10b). In this technique, the antibody (rather than the antigen) is immobilized on a microtiter well. A sample containing antigen is added and allowed to react with the immobilized antibody.After the well is washed, a second enzyme-linked antibody specific for a different epitope on the antigen is added and allowed to react with the bound antigen. After any free second antibody is removed by washing,substrate is added, and the colored reaction product is measured.
Another variation for measuring amounts of antigen is competitive ELISA (Figure 6-10c). In this technique, antibody is first incubated in solution with a sample containing antigen. The antigen-antibody mixture is then added to an antigencoated microtiter well. The more antigen present in the sample, the less free antibody will be available to bind to the antigen-coated well. Addition of an enzyme-conjugated secondary antibody (Ab2) specific for the isotype of the primary antibody can be used to determine the amount of primary antibody bound to the well as in an indirect ELISA. In the competitive assay, however, the higher the concentration of antigen in the original sample, the lower the absorbance.
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Measurement of light produced by chemiluminescence during certain chemical reactions provides a convenient and highly sensitive alternative to absorbance measurements in ELISA assays. In versions of the ELISA using chemiluminescence, a luxogenic (light-generating) substrate takes the place of the chromogenic substrate in conventional ELISA reactions.
For example, oxidation of the compound luminol by H2O2 and the enzyme horseradish peroxidase (HRP) produces light. The advantage of chemiluminescence assays over chromogenic ones is enhanced sensitivity. In general, the detection limit can be increased at least ten-fold by switching from a chromogenic to a luxogenic substrate, and with the addition of enhancing agents, more than 200-fold. In fact, under ideal conditions, as little as 5 1018 moles (5 attomoles) of target antigen have been detected.
A modification of the ELISA assay called the ELISPOT assay allows the quantitative determination of the number of cells in a population that are producing antibodies specific for a given antigen or an antigen for which one has a specific antibody (Figure 6-11). In this approach, the plates are coated with the antigen (capture antigen) recognized by the antibody
of interest or with the antibody (capture antibody) specific for the antigen whose production is being assayed. A suspension of the cell population under investigation is then added to the coated plates and incubated. The cells settle onto the surface of the plate, and secreted molecules reactive with the capture molecules are bound by the capture molecules in the vicinity of the secreting cells, producing a ring of antigen-antibody complexes around each cell that is producing the molecule of interest. The plate is then washed and an enzyme-linked antibody specific for the secreted antigen or specific for the species (e.g., goat anti-rabbit) of the secreted antibody is added and allowed to bind. Subsequent development of the assay by addition of a suitable chromogenic or chemiluminescence-producing substrate reveals the position of each antibody- or antigen-producing cell as a point of color or light.
ECL Assay Principles
Electrochemiluminescence (ECL) processes are known to occur with numerous molecules including compounds of ruthenium, osmium, rhenium or other elements. ECL is a process in which highly reactive species are generated from stable precursors at the surface of an electrode. These highly reactive species react with one another producing light. The development of ECL immunoassays is based on the use of a ruthenium chelate as the complex for the development of light. The chemiluminescent reactions that lead to the emission of light from the ruthenium complex are initiated electrically rather than chemically. This is achieved by applying a voltage to the immunological complexes (including the ruthenium complex) that are attached to Streptavidin - coated micro particles. Streptavidin, isolated from Streptomyces avidinii is preferred to avidin in this biotin- mediated immunoassay since it has an affinity for biotin comparable to that of avidin, is less basic and had no carbohydrate residues, thus limiting non - specific reactions with acidic groups
and lectins. The advantage of electrically initiating the chemiluminescent reaction is that the entire reaction can be precisely controlled.
The immunoprecipitation technique has the advantage of allowing the isolation of the antigen of interest for further analysis. It also provides a sensitive assay for the presence of a particular antigen in a given cell or tissue type.An extract produced by disruption of cells or tissues is mixed with an antibody against the antigen of interest in order to form an antigen-antibody complex that will precipitate. However, if the antigen concentration is low (often the case in cell and tissue extracts), the assembly of antigen-antibody complexes into precipitates can take hours, even days, and it is difficult to isolate the small amount of immunoprecipitate that forms. Fortunately, there are a number of ways to avoid these limitations. One is to attach the antibody to a solid support, such as a synthetic bead, which allows the antigen-antibody complex to be collected by centrifugation. Another is to add a secondary antibody specific for the primary antibody to bind the antigen-antibody complexes. If the secondary antibody is attached to a bead, the immune complexes can be collected by centrifugation. A particularly ingenious version of this procedure involves the coupling of the secondary antibody to magnetic beads. After the secondary antibody binds to the primary antibody, immunoprecipitates are collected by placing a magnet against the side of the tube
When used in conjunction with biosynthetic radioisotope labeling,immunoprecipitation can also be used to determinewhether a particular antigen is actually synthesized by a cell or tissue. Radiolabeling of proteins synthesized by cells of interest can be done by growing the cells in cell-culture medium containing one or more radiolabeled amino acids. Generally, the amino acids used for this application are those most resistant to metabolic modification, such as leucine, cysteine, or methionine. After growth in the radioactive medium, the cells are lysed and subjected to a primary antibody specific for the antigen of interest. The Ag-Ab complex is collected by immunoprecipitation, washed free of unincorporated radiolabeled amino acid and other impurities, and then analyzed. The complex can be counted in a scintillation counter to obtain a quantitative determination of the amount of the protein synthesized. Further analysis often involves disruption of the complex, usually by use of SDS and heat, so that the identity of the immunoprecipitated antigen can be confirmed by checking that its molecular weight is that expected for the antigen of interest. This is done by separation of the disrupted complex by SDS-PAGE and subsequent autoradiography to determine the position of the radiolabeled antigen on the gel.
In 1944, Albert Coons showed that antibodies could be labeled with molecules that have the property of fluorescence. Fluorescent molecules absorb light of one wavelength(excitation) and emit light of another wavelength (emission). If antibody molecules are tagged with a fluorescent dye, or fluorochrome, immune complexes containing these fluorescently labeled antibodies (FA) can be detected by colored light emission when excited by light of the appropriate wavelength. Antibody molecules bound to antigens in cells or tissue sections can similarly be visualized. The emitted light can be viewed with a fluorescence microscope,which is equipped with a UV light source. In this technique, known as immunofluorescence, fluorescent compounds such as fluorescein and rhodamine are in common use, but other highly fluorescent substances are also routinely used, such as phycoerythrin, an intensely colored and highly fluorescent pigment obtained from algae. These molecules can be conjugated to the Fc region of an antibody molecule without affecting the specificity of the antibody. Each of the fluorochromes below absorbs light at one wavelength and emits light at a longer wavelength:
Fluorescein, an organic dye that is the most widely used label for immunofluorescence procedures, absorbs blue light (490 nm) and emits an intense yellow-green fluorescence (517 nm).
Rhodamine, another organic dye, absorbs in the yellow-green range (515 nm) and emits a deep red fluorescence (546 nm). Because it emits fluorescence at a longer wavelength than fluorescein, it can be used in two-color immunofluorescence assays. An antibody specific to one determinant is labeled with fluorescein, and an antibody recognizing a different antigen is
labeled with rhodamine. The location of the fluorescein-tagged antibody will be visible by its yellowgreen color, easy to distinguish from the red color emitted where the rhodamine-tagged antibody has bound. By conjugating fluorescein to one antibody and rhodamine to another antibody, one can, for example, visualize simultaneously two different cell-membrane antigens on the same cell. Phycoerythrin is an efficient absorber of light (~30-fold greater than fluorescein) and a brilliant emitter of red fluorescence, stimulating its wide use as a label for immunofluorescence. Fluorescent-antibody staining of cell membrane molecules
or tissue sections can be direct or indirect (Figure 6-14). In direct staining, the specific antibody (the primary antibody) is directly conjugated with fluorescein; in indirect staining, the primary antibody is unlabeled and is detected with an additional fluorochrome-labeled reagent. A number of reagents have been developed for indirect staining.
The most common is a fluorochrome-labeled secondary antibody raised in one species against antibodies of another species, such as fluorescein-labeled goat anti-mouse immunoglobulin. Indirect immunofluorescence staining has two advantages over direct staining. First, the primary antibody does not need to be conjugated with a fluorochrome. Because the supply of primary antibody is often a limiting factor, indirect methods avoid the loss of antibody that usually occurs during the conjugation reaction. Second, indirect methods increase the sensitivity of staining because multiple molecules of the fluorochrome reagent bind to each primary antibody molecule, increasing the amount of light emitted at the location of each primary antibody molecule. Immunofluorescence has been applied to identify a number of subpopulations of lymphocytes, notably the CD4+ and CD8+ T-cell subpopulations. The technique is also suitable for identifying bacterial species, detecting Ag-Ab complexes in autoimmune disease, detecting complement components in tissues, and localizing hormones and other cellular products stained in situ. Indeed, a major application of the fluorescent-antibody technique is the localization of antigens in tissue sections or in subcellular compartments. Because it can be used to map the actual location of target antigens, fluorescence microscopy is a powerful tool for relating the molecular architecture of tissues and organs to their overall gross anatomy.
Flow Cytometry and Fluorescence
The fluorescent antibody techniques described are extremely valuable qualitative tools, but they do not give quantitative data. This shortcoming was remedied by development of the flow cytometer, which was designed to automate the analysis and separation of cells stained with fluorescent antibody. The flow cytometer uses a laser beam and light detector to count single intact cells in suspension. Every time a cell passes the laser beam, light is deflected from the detector, and this interruption of the laser signal is recorded. Those cells having a fluorescently tagged antibody bound to their cell surface antigens are excited by the laser and emit light that is recorded by a second detector system located at a right angle to the laser beam. The simplest form of the instrument counts each cell as it passes the laser beam and records the level of fluorescence the cell emits; an attached computer generates plots of the number of cells as the ordinate and their fluorescence intensity as the abscissa. More sophisticated versions of the instrument are capable of sorting populations of cells into different containers according to their fluorescence profile. Use of the instrument to determine which and how many members of a cell population bind fluorescently labeled antibodies is called analysis; use of the instrument to place cells having different patterns of reactivity into different containers is called cell sorting. The flow cytometer has multiple applications to clinical and research problems. A common clinical use is to determine the kind and number of white blood cells in blood samples. By treating appropriately processed blood samples with a fluorescently labeled antibody and performing flow cytometric analysis, one can obtain the following information:
How many cells express the target antigen as an absolute number and also as a percentage of cells passing the beam. For example, if one uses a fluorescent antibody specific for an antigen present on all T cells, it would be possible to determine the percentage of T cells in the total white blood cell population. Then, using the cell-sorting capabilities of the flow cytometer, it would be possible to isolate the T-cell fraction of the leukocyte population.
The distribution of cells in a sample population according to antigen densities as determined by fluorescence intensity. It is thus possible to obtain a measure of the distribution of antigen density within the population of cells that possess the antigen. This is a powerful feature of the instrument, since the same type of cell may express different levels of antigen depending
upon its developmental or physiological state.
The size of cells. This information is derived from analysis of the light-scattering properties of members of the cell population under examination.
Flow cytometry also makes it possible to analyze cell populations that have been labeled with two or even three different fluorescent antibodies. For example, if a blood sample is reacted with a fluorescein-tagged antibody specific for T cells, and also with a phycoerythrin-tagged antibody specific for B cells, the percentages of B and T cells may be determined simultaneously with a single analysis.Numerous variations of such "two-color" analyses are routinely carried out, and "three-color" experiments are common.Aided by appropriate software, highly sophisticated versions of the flow cytometer can even perform "five-color" analyses. Flow cytometry now occupies a key position in immunology and cell biology, and it has become an indispensable clinical tool as well. In many medical centers, the flow cytometer is one of the essential tools for the detection and classification of leukemias (see the Clinical Focus). The choice of treatment for leukemia depends heavily on the cell types involved, making precise identification of the neoplastic cells an essential part of clinical practice. Likewise, the rapid measurement of T-cell subpopulations, an important prognostic indicator in AIDS, is routinely done by flowcytometric analysis. In this procedure, labeled monoclonal antibodies against the major T-cell subtypes bearing the CD4 and CD8 antigens are used to determine their ratios in the patient's blood. When the number of CD4 T cells falls
below a certain level, the patient is at high risk for opportunistic infections.