Methods and Concepts in the Life Sciences/Immunoassays
The western blot (sometimes called protein immunoblot) is a widely used analytical technique used to detect specific proteins in a sample. It uses gel electrophoresis to separate native proteins by 3-D structure or denatured proteins by the length of the polypeptide. The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are stained with antibodies specific to the target protein. The gel electrophoresis step is included in western blot analysis to resolve the issue of the cross-reactivity of antibodies. A number of search engines, such as CiteAb, are available that can help researchers find suitable antibodies for use in western blotting.
The name western blot is a play on the name Southern blot, a technique for DNA detection developed earlier by Edwin Southern. Detection of RNA is termed northern blot.
Fehler: Referenz nicht gefunden Fehler: Referenz nicht gefundenThe proteins of the sample are separated using gel electrophoresis. Separation of proteins may be by isoelectric point (pI), molecular weight, electric charge, or a combination of these factors. SDS-PAGE is by far the most common type of gel electrophoresis. It is also possible to use a two-dimensional gel electrophoresis.
In order to make the proteins accessible to antibody detection, they are moved from within the gel onto a membrane made of nitrocellulose or polyvinylidene difluoride (PVDF). The primary method for transferring the proteins is called electroblotting and uses an electric current to pull proteins from the gel into the membrane. An older method of transfer involves placing a membrane on top of the gel, and a stack of filter papers on top of that. The entire stack is placed in a buffer solution which moves up the paper by capillary action, bringing the proteins with it. In practice this method is not used anymore as it takes too much time. As a result of either blotting process, the proteins are exposed on a thin surface layer for detection (see below). Both varieties of membrane are chosen for their non-specific protein binding properties (i.e. binds all proteins equally well). Protein binding is based upon hydrophobic interactions, as well as charged interactions between the membrane and protein. Nitrocellulose membranes are cheaper than PVDF, but are far more fragile and do not stand up well to repeated probings.
The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie Brilliant Blue or Ponceau S dyes. Ponceau S is the more common of the two, due to its higher sensitivity and water solubility, the latter making it easier to subsequently destain and probe the membrane, as described below.
Since the membrane has been chosen for its ability to bind protein, steps must be taken to prevent the interactions between the membrane and the antibody used for detection of the target protein. Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein - typically 3-5% Bovine serum albumin (BSA) or non-fat dry milk (both are inexpensive) in Tris-Buffered Saline (TBS) or I-Block, with a minute percentage (0.1%) of detergent such as Tween 20 or Triton X-100. The protein in the dilute solution attaches to the membrane in all places where the target proteins have not attached. Thus, when the antibody is added, there is no room on the membrane for it to attach other than on the binding sites of the specific target protein. This reduces "noise" in the final product of the western blot, leading to clearer results, and eliminates false positives.
During the detection process the membrane is "probed" for the protein of interest with a modified antibody which is linked to a reporter enzyme. When exposed to an appropriate substrate, this enzyme drives a colourimetric reaction and produces a color. For a variety of reasons, this traditionally takes place in a two-step process, although there are now one-step detection methods available for certain applications.
In the two-step process, a dilute solution of primary antibody (generally between 0.5 and 5 micrograms/mL) is incubated with the membrane under gentle agitation. Typically, the solution is composed of buffered saline solution with a small percentage of detergent, and sometimes with powdered milk or BSA. The antibody solution and the membrane can be sealed and incubated together for anywhere from 30 minutes to overnight. It can also be incubated at different temperatures, with higher temperatures being associated with more binding, both specific (to the target protein, the "signal") and non-specific ("noise").
After rinsing the membrane to remove unbound primary antibody, the membrane is exposed to another antibody, directed at a species-specific portion of the primary antibody. Antibodies come from animal sources (or animal sourced hybridoma cultures). An anti-mouse secondary will bind to almost any mouse-sourced primary antibody, which allows some cost savings by allowing an entire lab to share a single source of mass-produced antibody, and provides far more consistent results. The secondary antibody is usually linked to biotin or to a reporter enzyme such as alkaline phosphatase or horseradish peroxidase. This means that several secondary antibodies will bind to one primary antibody and enhance the signal.
Most commonly, a horseradish peroxidase-linked secondary is used to cleave a chemiluminescent agent, and the reaction product produces luminescence in proportion to the amount of protein. A sensitive sheet of photographic film is placed against the membrane, and exposure to the light from the reaction creates an image of the antibodies bound to the blot. A cheaper but less sensitive approach utilizes a 4-chloronaphthol stain with 1% hydrogen peroxide; reaction of peroxide radicals with 4-chloronaphthol produces a dark purple stain that can be photographed without using specialized photographic film.
Another method of secondary antibody detection utilizes a near-infrared (NIR) fluorophore-linked antibody. Light produced from the excitation of a fluorescent dye is static, making fluorescent detection a more precise and accurate measure of the difference in signal produced by labeled antibodies bound to proteins on a western blot. Proteins can be accurately quantified because the signal generated by the different amounts of proteins on the membranes is measured in a static state, as compared to chemiluminescence, in which light is measured in a dynamic state.
A third alternative is to use a radioactive label rather than an enzyme coupled to the secondary antibody, such as labeling an antibody-binding protein like Staphylococcus Protein A or Streptavidin with a radioactive isotope of iodine. Since other methods are safer, quicker, and cheaper, this method is now rarely used; however, an advantage of this approach is the sensitivity of auto-radiography-based imaging, which enables highly accurate protein quantification when combined with optical software (e.g. Optiquant).
Historically, the probing process was performed in two steps because of the relative ease of producing primary and secondary antibodies in separate processes. This gives researchers and corporations huge advantages in terms of flexibility, and adds an amplification step to the detection process. Given the advent of high-throughput protein analysis and lower limits of detection, however, there has been interest in developing one-step probing systems that would allow the process to occur faster and with fewer consumables. This requires a probe antibody which both recognizes the protein of interest and contains a detectable label, probes which are often available for known protein tags. The primary probe is incubated with the membrane in a manner similar to that for the primary antibody in a two-step process, and then is ready for direct detection after a series of wash steps.
After the unbound probes are washed away, the western blot is ready for detection of the probes that are labeled and bound to the protein of interest. In practical terms, not all westerns reveal protein only at one band in a membrane. Size approximations are taken by comparing the stained bands to that of the marker or ladder loaded during electrophoresis. The process is commonly repeated for a structural protein, such as actin or tubulin, that should not change between samples. The amount of target protein is normalized to the structural protein to control between groups. A superior strategy is the normalization to the total protein visualized with trichloroethanol or epicocconone. This practice ensures correction for the amount of total protein on the membrane in case of errors or incomplete transfers.
The colorimetric detection method depends on incubation of the western blot with a substrate that reacts with the reporter enzyme (such as peroxidase) that is bound to the secondary antibody. This converts the soluble dye into an insoluble form of a different color that precipitates next to the enzyme and thereby stains the membrane. Development of the blot is then stopped by washing away the soluble dye. Protein levels are evaluated through densitometry (how intense the stain is) or spectrophotometry.
Chemiluminescent detection methods depend on incubation of the western blot with a substrate that will luminesce when exposed to the reporter on the secondary antibody. The light is then detected by CCD cameras which capture a digital image of the western blot or photographic film. The use of film for western blot detection is slowly disappearing because of non linearity of the image (non accurate quantification). The image is analysed by densitometry, which evaluates the relative amount of protein staining and quantifies the results in terms of optical density. Newer software allows further data analysis such as molecular weight analysis if appropriate standards are used. =
Radioactive labels do not require enzyme substrates, but rather allow the placement of medical X-ray film directly against the western blot, which develops as it is exposed to the label and creates dark regions which correspond to the protein bands of interest. The importance of radioactive detections methods is declining due to its hazardous radiation, because it is very expensive, health and safety risks are high, and ECL (enhanced chemiluminescence) provides a useful alternative.
Fluorescently labeled probes are excited by light and the emission of the excitation is then detected by a photosensor such as a CCD camera equipped with appropriate emission filters which captures a digital image of the western blot and allows further data analysis such as molecular weight analysis and a quantitative western blot analysis. Fluorescence is considered to be one of the best methods for quantification, but is less sensitive than chemiluminescence.
The enzyme-linked immunosorbent assay (ELISA) is a test that uses antibodies and color change to detect a substance. The ELISA has been used as a diagnostic tool in medicine and plant pathology, as well as a quality-control check in various industries.
Performing an ELISA involves at least one antibody with specificity for substance of interest. The sample with an unknown amount of this antigen is immobilized on a solid support (usually a polystyrene microtiter plate) either non-specifically (via adsorption to the surface) or specifically (via capture by another antibody specific to the same antigen, in a "sandwich" ELISA). After the antigen is immobilized, the detection antibody is added, forming a complex with the antigen. The detection antibody can be covalently linked to an enzyme, or can itself be detected by a secondary antibody that is linked to an enzyme. Between each step, the plate is typically washed with a mild detergent solution to remove any proteins or antibodies that are non-specifically bound. After the final wash step, the plate is developed by adding an enzymatic substrate to produce a visible signal, which indicates the quantity of antigen in the sample.
The use of a secondary antibody avoids the expensive process of creating enzyme-linked antibodies for every antigen one might want to detect. By using an enzyme-linked antibody that binds the Fc region of other antibodies, this same enzyme-linked antibody can be used in a variety of situations. Enzymes used in ELISAs include horseradish peroxidase (HRP), alkaline phosphatase (AP) or glucose oxidase. These enzymes allow for detection because they produce an observable color change in the presence of certain reagents. In some cases these enzymes are exposed to reagents which cause them to produce light or Chemiluminescence. Enzymes are used because they act as amplifiers, producing a detectable signal even if only few enzyme-linked antibodies remain bound.
A direct ELISA consists of the following steps:
- A buffered solution of the antigen to be tested for is added to each well of a microtiter plate, where it is given time to adhere to the plastic through charge interactions.
- A solution of nonreacting protein, such as bovine serum albumin or casein, is added to well in order to cover any plastic surface in the well which remains uncoated by the antigen.
- The primary antibody is added, which binds specifically to the test antigen coating the well. This primary antibody could also be in the serum of a donor to be tested for reactivity towards the antigen.
- A secondary antibody is added, which will bind the primary antibody. This secondary antibody often has an enzyme attached to it, which has a negligible effect on the binding properties of the antibody. In other cases, as shown in figure 5, the primary antibody itself is conjugated to the enzyme.
- A substrate for this enzyme is then added. Often, this substrate changes color upon reaction with the enzyme. The color change shows the secondary antibody has bound to primary antibody, which strongly implies the donor has had an immune reaction to the test antigen. This can be helpful in a clinical setting, and in research.
- The higher the concentration of the primary antibody present in the serum, the stronger the color change. Often, a spectrometer is used to give quantitative values for color strength.
The use and meaning of the names "direct ELISA" and "indirect ELISA" differs in the literature depending on the context of the experiment. When the presence of an antigen is analyzed, the name "direct ELISA" refers to an ELISA in which only a labelled primary antibody is used, and the term "indirect ELISA" refers to an ELISA in which the antigen is bound by the primary antibody which then is detected by a labelled secondary antibody. In the latter case a sandwich ELISA is clearly distinct from an indirect ELISA. When the 'primary' antibody is of interest, e.g. in the case of immunization analyses, this antibody is directly detected by the secondary antibody and the term "direct ELISA" applies to a setting with two antibodies.
A major disadvantage of the direct ELISA is the method of antigen immobilization is not specific; when serum is used as the source of test antigen, all proteins in the sample may stick to the microtiter plate well, so small concentrations of analyte in serum must compete with other serum proteins when binding to the well surface. The sandwich ELISA provides a solution to this problem by using a capture antibody specific for the test antigen to pull it out of the serum's molecular mixture. The binding of this capture antibody to the surface is the first step in a sandwich ELISA. After blocking nonspecific binding sites on the surface the sample is applied and, if present, the antigen is bound by the capture antibody. The following procedure is identical to direct ELISA.
Use of the purified specific antibody to attach the antigen to the plastic eliminates a need to purify the antigen from complicated mixtures before the measurement, simplifying the assay, and increasing the specificity and the sensitivity of the assay.
A third use of ELISA is through competitive binding. The steps for this ELISA are somewhat different from the first two examples:
- Unlabeled antibody is added to the sample. If the sample contains the antigen, it is bound by the antibody.
- These antibody/antigen complexes are then added to an antigen-coated well.
- The plate is washed, so unbound antibody is removed. The more antigen in the sample, the more Ag-Ab complexes are formed and so there are less unbound antibodies available to bind to the antigen in the well.
- The secondary antibody, specific to the primary antibody, is added. This second antibody is coupled to an enzyme.
- A substrate is added, and remaining enzymes elicit a chromogenic or fluorescent signal.
- The reaction is stopped to prevent eventual saturation of the signal.
Some competitive ELISA kits include enzyme-linked antigen rather than enzyme-linked antibody. The labeled antigen competes for primary antibody binding sites with the sample antigen (unlabeled). The less antigen in the sample, the more labeled antigen is retained in the well and the stronger the signal.
The Enzyme-Linked ImmunoSpot (ELISPOT) assay is a widely used method for monitoring cellular immune responses in humans and animals, and has found clinical applications in the diagnosis of Tuberculosis and the monitoring of graft tolerance or rejection in transplant patients. The ELISPOT technique has proven to be among the most useful means available for monitoring Cell-Mediated Immunity, due to its sensitive and accurate detection of rare antigen-specific T cells (or B cells) and its ability to visualize single positive cells within a population of Peripheral Blood Mononuclear Cells (PBMC).
Simply put, at appropriate conditions the ELISPOT assay allows visualization of the secretory product(s) of individual activated or responding cells. Each spot that develops in the assay represents a single reactive cell. Thus, the ELISPOT assay provides both qualitative (regarding the specific cytokine or other secreted immune molecule) and quantitative (the frequency of responding cells within the test population) information.
In an ELISPOT assay, the membrane surfaces in a 96-well PVDF-membrane microtiter plate are coated with capture antibody that binds a specific epitope of the cytokine being assayed. During the cell incubation and stimulation step, PBMC are seeded into the wells of the plate along with the antigen, and form a monolayer on the membrane surface of the well. As the antigen-specific cells are activated, they release the cytokine, which is captured directly on the membrane surface by the immobilized antibody, before it has a chance to diffuse into the culture supernatant, or to be degraded by proteases and bound by receptors on bystander cells. Subsequent detection steps visualize the immobilized cytokine as an ImmunoSpot; essentially the secretory footprint of the activated cell.
The practical limits of detection for ELISPOT are dependent generally on the number of cells seeded in an assay well. Typically, 200,000 - 400,000 PBMC will be used per well for an assay, but up to one million cells are commonly used for detection of rare events. ELISPOT is capable of detecting a single antigen positive cell within this population.
The ELISPOT assays employ a technique very similar to the sandwich ELISA technique. Either a monoclonal (preferred for greater specificity) or polyclonal capture antibody is coated aseptically onto a PVDF (polyvinylidene fluoride) -backed microplate. The plate is blocked, usually with a serum protein that is non-reactive with any of the antibodies in the assay. After this, cells of interest are plated out at varying densities, along with antigen or mitogen, and then placed in a humidified 37°C CO2 incubator for a specified period of time.
Cytokine (or other cell product of interest) secreted by activated cells is captured locally by the coated antibody on the high surface area PVDF membrane. After washing the wells to remove cells, debris, and media components, a biotinylated polyclonal antibody specific for the chosen analyte is added to the wells. This antibody is reactive with a distinct epitope of the target cytokine and thus is employed to detect the captured cytokine. Following a wash to remove any unbound biotinylated antibody, the detected cytokine is then visualized using streptavidin conjugated to an enzyme — horseradish peroxidase (HRP) or alkaline phosphatase (AP) — and a precipitating substrate (e.g., AEC, BCIP/NBT). The colored end product (a spot, usually red (for HRP) or a blackish blue (for AP)) typically represents an individual cytokine-producing cell. The spots can be counted manually (e.g., with a dissecting microscope) or using an automated reader to capture the microwell images and to analyze spot number and size.
Lateral flow test
Lateral flow tests, also known as lateral flow immunochromatographic assays, are simple devices intended to detect the presence (or absence) of a target analyte in a sample without the need for specialized and costly equipment, though many lab-based applications exist that are supported by reading equipment. Typically, these tests are used for medical diagnostics. A widely spread and well known application is the home pregnancy test.
The technology is based on a series of capillary beds, such as pieces of porous paper or sintered polymer. Each of these elements has the capacity to transport fluid (e.g., urine) spontaneously. The first element (the sample pad) acts as a sponge and holds an excess of sample fluid. Once soaked, the fluid migrates to the second element (conjugate pad) in which the manufacturer has stored the so-called conjugate, a dried format of bio-active particles in a salt-sugar matrix that contains everything to guarantee an optimized chemical reaction between the target molecule (e.g., an antigen) and its chemical partner (e.g., antibody) that has been immobilized on the particle's surface. While the sample fluid dissolves the salt-sugar matrix, it also dissolves the particles and in one combined transport action the sample and conjugate mix while flowing through the porous structure. In this way, the analyte binds to the particles while migrating further through the third capillary bed. This material has one or more areas (often called stripes) where a third molecule has been immobilized. By the time the sample-conjugate mix reaches these strips, analyte has been bound on the particle and the third 'capture' molecule binds the complex. After a while, when more and more fluid has passed the stripes, particles accumulate and the stripe-area changes color. Typically, there are at least two stripes: one (the control) that captures any particle and thereby shows that reaction conditions and technology worked fine, and a second that contains a specific capture molecule and only captures those particles onto which an analyte molecule has been immobilized. After passing these reaction zones the fluid enters the final porous material, the wick, that simply acts as a waste container. Lateral flow tests can operate as either competitive or sandwich assays.