Selected analytical laboratory techniques
The immunological reaction between antigen and antibody forms the basis of a pattern of laboratory diagnostic tests. The binding of antigen (Ag) to antibody (Ab) in aqueous solution involves a specific, reversible association to form an antigen-antibody complex (immune complex) while maintaining their structure.
The bipolar centers of the association reaction of antigen and antibody are the binding sites on the variable domain of the immunoglobulin molecule and the antigenic determinant (epitope) of the antigen.
The epitope is the antigenic determinant or the portion of the antigen surface that is recognized by the binding site on the antibody. It consists of approximately 30 amino acids, although X-ray structure analysis has indicated that only up to 17 amino acids at most are involved in antibody binding .
Antibodies are immunoglobulins. They are Y-shaped glycoproteins that consist of four polypeptide chains (two identical heavy (H) chains and two identical light (L) chains) linked by disulfide bonds. Pepsin cleaves the F(ab)2 fragment from the Fc region of the antibody and papain treatment produces Fab fragments.
The H and L chains of the immunoglobulin molecule contain subunits (also known as domains), some of which are identical in all antibodies of the same isotype (constant domains) and some of which differ (variable domains). The antigen binding site includes three variable domains of the L chain and three variable domains of the H chain on each arm of the Y-shaped molecule: the complementarity determining regions (CDRs). The specific chemistry, nature, and structure of the epitopes bound are determined by these CDRs /, /. The site on the antibody to which the epitope binds is called the paratope. The paratope forms part of the surface structure of the antigen binding site known as the idiotype. Paratopes are made up of six highly accessible loops of hyper variable sequence, totaling 50–60 amino acid residues.
- The complementarity of paratope and epitope surfaces which are in contact, so that depressions in one are by and large filled by protrusions from the other
- The complementary in the physical and chemical properties of the interfacing surfaces. The contacts involve hydrogen bonds, and van der Waal’s interactions (hydrogen bonds form between polar groups and oppositely charged side chains form ion pairs).
The idiotype describes the sum of all idiotopes on an antibody.
Anti-idiotypic antibodies are directed against the hyper variable region of the antigen binding site of an immunoglobulin molecule.
Anti-isotypic antibodies are directed against a constant region of the immunoglobulin molecule.
The major components of an immunoassay are the antigen and the antibody. In a typical immunoassay:
- The antibody is used as a reagent to detect the antigen (substance of interest, e.g. parathyroid hormone)
- The antigen is used to detect an antibody of interest (e.g., anti-Rubella antibody).
To produce specific antibodies, the antigen must either be an immunogen or be altered in such a way that it acts as an immunogen.
An immunogen is a substance (protein or a substance coupled to a carrier) that induces an immune response when introduced in a foreign body. The immunogen used in the immunoassay is either identical to the analyte or has the same characteristic features.
There are some differences between immunogens and antigens. An antigen binds to a specific antibody. While all immunogens are antigens, not all antigens are immunogens. The antigens used as reagents in an immunoassay are purified immunogens, prepared in such a way that the destruction of partial antigens or the physical antigen structure does not affect the specificity of the immunoassay.
The function of the antibody is to ensure that the immunoassay is specific for the analyte (immunogen) and to enable the analyte to be quantified. The antibodies used in immunoassays may be polyclonal, monoclonal, mixed monoclonal, or polyclonal-monoclonal. In general, if the antibody sensitivity and specificity are high, the detection limit and analytical specificity of the immunoassay is guaranteed to meet requirements. Occasionally, however, high affinity may be associated with decreased specificity; for this reason, the criteria required for a reliable analysis must be specified precisely. Polyclonal and monoclonal antibodies are used in immunoassays .
Polyclonal antibodies are produced by immunizing animals with the immunogen of interest. Usually, this yields an antiserum that is based on polyclonal antibody synthesis and that contains a pattern of antibodies of varying analytic sensitivity and specificity, avidity, valency, and binding kinetics directed against different epitopes on the immunogen. The antibody with optimal immunoassay characteristics is selected.
Monoclonal antibodies are produced by fusing antibody secreting B cells from the spleen of an antigen sensitized mouse with myeloma cells (in vitro hybridization), followed by cloning and cultivation of the hybrid cells in cell culture. Ideally, the monoclonal antibody produced is directed against one epitope of the antigen. Characteristics are identical specificity, avidity, valency, and binding kinetics. Monoclonal antibodies belong to a single immunoglobulin (Ig) class and type and have identical specificity. All molecules have identical physicochemical properties. Monoclonal antibodies exhibit very limited structural diversity and are homogeneous in comparison to polyclonal antibodies.
Depending on the intended use, monoclonal antibodies do not always offer advantages over polyclonal antibodies purified using affinity chromatography. For example, monoclonal antibodies usually have lower affinity and avidity than polyclonal antibodies.
The ability of monoclonal antibodies to recognize only one epitope can be a disadvantage:
- For species-wide or genus-wide recognition of antigens of infectious pathogens
- In immunoprecipitation methods with low epitope density of the antigen
- If the antigen is also expressed by other pathogens, for example, this can result in a lack of specificity and cross reactivity.
The manufacturers of immunoassays tackle this problem by using a cocktail of monoclonal antibodies of differing epitope specificities.
Monoclonal antibody production
Antigen sensitized B lymphocytes with the information to produce specific antibodies are fused with myeloma plasma cells. The myeloma cells are generated by injecting a mouse with mineral oil to induce myeloma formation. This results in an immortal plasma cell line. As an immunoglobulin factory, the fused cell, also referred to as a hybrid cell, is capable of producing antibodies of one class and one type with identical characteristics /, /.
The information for the antibody production is brought into the hybrid cells by sensitized B cells from the spleens of mice that have been immunized with the relevant antigen. Antigen sensitized B cells are only able to survive a few days in vitro.
The hybridized mouse plasma cells from myeloma cell lines contribute immortality as well as immunoglobulin synthesis.
Antigen sensitized B cells and plasma cannot survive for more than a few days. After fusion, the cells are placed into HAT (hypoxanthine, aminopterin, and thymidine) medium. There, the quickly growing non fused antigen sensitized B cells and plasma cells die off while the hybrid cells are maintained.
The non fused myeloma cells die as a result of the inhibition of purine synthesis by the folic acid antagonist aminopterin. Mutant myeloma cells instead of normal plasma cells are fused with the antigen sensitized B cells. Because the mutant myeloma cells are deficient in hypoxanthine-guanine-phosphoribosyl-transferase (HGPRT) and thymidine kinase, they cannot synthesize purines from thymidine and hypoxanthine in the presence of aminopterin.
The hybrid cells can synthesize purines, however, since the antigen sensitized B cells provide an intact HGPRT alternative pathway.
After the hybrid cells have been cultivated for a number of weeks, they are screened for immunoglobulin production and subcultured so that, in theory, a cell clone can be produced from a single hybrid cell (cloning step).
As the last step, the different clones are checked for their ability to produce an antibody with optimal characteristics and the clone that produces this antibody is selected for large scale antibody production. This takes place either in in-vitro mass culture or in the ascites of laboratory animals. One hybrid cell produces approximately 50 ng of immunoglobulin daily.
The simplest immune reaction is that between a single antigen (epitope) and its corresponding binding site (paratope) on the Fab fragment of a monomeric antibody molecule. The strength of the bond between the two reactants (primary interaction) is the net result of the attracting and repelling forces exerted by each on the other and is referred to as the antibody affinity or intrinsic affinity. Because this is a thermodynamic reaction, it is described by an equilibrium constant (K):
K = AgAb/Ag × Ab
High affinity antibodies dissociate from the AgAb complex with difficulty and require a high level of energy to do so.
Ideally, antibody affinity can only be measured between monoclonal antibodies and epitopes. If complex antigens and polyclonal antisera are used, it is only possible to determine the mean antibody avidity.
An antibody is a specific immunoglobulin produced by a plasma cell (differentiated B cell) in response to contact between the B cell and an immunogen. The immunoglobulin is synthesized by a clone of plasma cells.
Antigens stimulate the production of antibodies and bind to these antibodies.
An assay is a quantitative test used to determine the quantity, activity, or strength of a component or characteristic.
Avidity represents the net affinity of all binding sites of all antibodies in the antiserum under specified physicochemical reaction conditions.
This is the capacity of a receptor such as an IgM antibody to bind to a ligand such as an antigen.
This is the quantitatively measured value of an analyte that is used to determine whether a result lies above or below a clinical or analytic threshold value.
The epitope is the chemical group on an antigen to which an antibody can bind.
A fluorophore is a substance that emits light when it is supplied with electromagnetic radiation.
Heterophilic antibodies react with antigens from another species of animal. For instance, the antibodies to viral antigens that are produced in Epstein-Barr viral infections also cross react with epitopes on the red blood cells of sheep and thus cause hemagglutination (Paul-Bunnell reaction).
This is a property of an antiserum that reacts with different antigens.
A heterogeneous assay requires the physical separation of unbound antigen from bound immune complex antigen before the antigen can be quantified.
In this type of immunoassay, the analyte (sample) and the immunochemical reagents (antibody or antibody conjugate) are combined in the incubation mixture and a washing step is not required before the bound fraction is measured. A prerequisite is that the analyte generates a measurable dose-response signal after antibody binding that differentiates between bound and unbound analyte.
Immune complex (antigen-antibody complex)
Antigens react with antibodies in aqueous solution to form immune complexes.
The formation of immune complexes declines:
- As pH decreases (pH < 7)
- As ionic strength increases (very rapid reaction in an ion-poor solution)
- As temperature increases.
The size of immune complexes and their solubility depends on the ratio of the number of antigen molecules to the number of available antibody binding sites in the reagent mixture. In antibody or antigen excess, immune complexes are soluble and smaller than if there is antigen-antibody equivalence. When the equivalence zone is reached, large, insoluble, precipitating immune complexes result.
The immunonephelometric and immunoturbidimetric methods for the determination of unknown antigen concentrations in free solution employ a moderate antibody excess. Under such conditions, immune complexes form that are still soluble and up to 0.5 μm in size, which corresponds to a molecular mass of up to 100,000 kDa. Such a complex contains approximately 200 antigen and 200 antibody molecules.
Immunopotency is a characteristic of an antibody in an immune reaction. The potency of an antibody depends on its avidity and its concentration. The potency of an antigen in an immune reaction depends on its concentration.
A calibrator is a solution adjusted to a standardized reference material and used to calibrate an assay.
In a competitive immunoassay, the labeled and unlabeled analyte compete for a binding protein or receptor.
Cross reactivity describes the reaction of an antibody with an antigen other than the antigen that elicited its formation. It occurs due to the presence of shared, identical, or similar antigenic determinants.
A label is a signal generating substance used to mark an antigen or antibody to make an immune reaction visible. Radioactivity, luminophores, fluorophores, and enzymes are used as labels.
The ligand is a substance or part of a substance that binds to a receptor, antibody, or other binding protein.
A luminophore is a molecule that emits luminescence in the form of photons following stimulation.
Post zone effect
The post zone effect refers to the increased solubility of immune complexes as the result of an antigen concentration that is significantly in excess of the antibody concentration.
The prozone effect is the result of a suboptimal antigen-antibody reaction. It occurs due to an excess of antigen or antibody or because one of the reaction partners is incomplete or blocks an optimal reaction.
The specificity describes the quality of an antiserum to react with defined antigens. In the case of an immunoassay, the specificity is a criterion of the extent to which the assay responds only to a specified analyte and not to other substances present in the sample.
The specific activity is a measure of events per unit of mass or time.
A tracer is a labeled analyte for measuring the dose-response characteristics of an immunoassay.
Antigen or antibody determination in vitro is only possible if the antigen-antibody reaction can be visualized or is measurable. The selection of the method of determination depends on the characteristics of the antigen (size, number, and structure of the antigenic determinants), the characteristics of the corresponding antibody (avidity, specificity), and the concentration of the analyte to be determined.
Antigens or antibodies can be measured according to the following principles:
- Direct detection
- Indirect detection
- Detection based on the labeling of antigen or antibody.
Immunochemical techniques for detection and quantification of an unknown antibody employ a defined amount of antigen while methods for the detection of an unknown antigen concentration use a defined concentration of antibody. The critical factors that determine whether the immune complexes formed can be detected in a soluble form or as precipitate include an optimal ratio of the antigen concentration to the antibody concentration. For the precipitin reaction the model of Heidelberger and Kendall describes the results observed when increasing concentrations of antigens are mixed with a constant concentration of antibody ().
In antibody excess, small, soluble immune complexes are formed. Their concentration, measured as a scattered light signal (e.g., immunonephelometry) or in the form of absorption (e.g., immunoturbidimetry), is proportional to the antigen concentration.
With an increase in the antigen concentration, the equivalence is reached and large, insoluble, precipitating immune complexes result. In all immunoprecipitation methods such as radial immunodiffusion or immunofixation electrophoresis, the antigen-antibody systems are adjusted in such a manner that the tests are run within the equivalence zone.
In the case of antigen excess, mainly small soluble immune complexes are present. Their concentration increases with increasing antigen concentration and the measurement signal (scattered light, absorption) or the precipitate quantity declines, mimicking too low an antigen concentration. This is the post zone effect.
In order to obtain a signal that is proportional to the antigen concentration, the specimen must be diluted to such a degree that the antigen concentration falls within the range of the antibody excess. This corresponds to the ascending slope of the Heidelberger and Kendall curve ().
Precipitin techniques (e.g., radial immunodiffusion, immunoelectrophoresis) and techniques involving measurements in a liquid phase such as immunonephelometry and immunoturbidimetry are aimed at the direct detection of antigens or antibodies.
Precipitin reactions that allow the direct visualization of the immunoprecipitate have a low detection limit and only the detection of high concentrations of antigen is possible. However, if the antigen or antibody is bound to a solid phase, thus creating multi valency, the antigen-antibody reaction is visible in the form of an agglutination ().
Methods that employ a solid phase for antigens or antibodies include the following methods: latex agglutination, indirect (passive) hemagglutination, the hemagglutination inhibition test, and the complement fixation test.
Labeled immunochemical assays
These assays are used for the detection or quantification of antigens or antibodies in low concentrations. Antigens or antibodies are labeled with:
- A fluorophore (immunofluorescence assay)
- Radioactivity (radioimmunoassay)
- An enzyme (enzyme immunoassay)
- A chemiluminescence label (luminescence immunoassay).
The hemagglutination test for the determination of blood groups and the bacterial agglutination test (Widal reaction) are direct agglutination assays.
Blood group determination
Agglutination of bacteria
Detection of antibodies (Widal reaction)
Principle: suspensions of inactivated bacteria are used as antigen and incubated with dilutions of the patient’s serum. The occurrence of agglutination indicates the presence of the corresponding antibody in the patient’s serum.
Detection of antigens (Gruber reaction)
Principle: for the classification and typing of bacteria, cultures are incubated with appropriate class and type-specific antisera.
This method is based on the principle that in the presence of a specified amount of antibody, the antigen is diluted by diffusion in a semisolid medium (e.g., agarose gel) until antigen-antibody equivalence is reached, at which maximum precipitate occurs.
Principle: the test is performed using agar plates that contain corresponding antibodies to the antigen of interest. The sample is applied to holes in the gel. The antigen in the sample diffuses radially into the gel /, /.
In the measurement according to Mancini, diffusion is allowed to occur until the antigen has completely precipitated. According to the Heidelberger and Kendall curve, small, soluble immune complexes initially form close to the application site. With increasing diffusion from the site of application, larger, insoluble complexes are formed. When excess antigen is concerned, the point of equivalence is reached and the diffusion of the antigen terminated due to complete precipitation. The antigen concentration is proportional to the second power of the diameter of the precipitate ring.
In the measurement according to Fahey, the concentration dependent diffusion speed of the antigen is used for the assessment. Therefore, in comparison to the Mancini method, the reaction time is shortened. The antigen concentration is proportional to the diameter of the precipitate ring.
In both methods, the antigen concentration is determined using a calibration curve based on calibrators that are run simultaneously with the patient’s sample or directly by means of factors in the case of standardized plates.
Principle: two procedures are combined: serum protein electrophoresis and immunoprecipitation /, /. Agarose gel or cellulose acetate membranes are used as the support medium while customary electrophoresis buffer at a pH of about 8.5 is employed as the separation medium. Following electrophoretic separation of the patient’s sample and a reference sample, a trough is cut in the gel in the direction of separation and filled with an antiserum against the protein of interest. The electrophoresis plate is placed into a humid chamber for 16–24 h. During this time simultaneous diffusion of the antiserum from the trough and the antigens from the separated sample results in the formation of precipitin arcs. According to the position, shape, and strength of the precipitin arcs, individual proteins can be identified and their concentration can be crudely estimated.
Immunoelectrophoresis is indicated if serum monoclonal immunoglobulin or immunoglobulin fragments are suspected. They belong to one immunoglobulin (Ig) class and/or one Ig type and are present in excess. Following their electrophoretic separation, polyclonal Ig are homogeneously distributed within the γ-globulin fraction and the precipitate arc that forms during the diffusion phase has an evenly curved shape. Monoclonal antibodies or antibody fragments form a local enhancement (M gradient) within the γ-globulin arc. In the diffusion phase, the equivalence zone shifts towards the antiserum trough and the precipitate arc is bowed and/or thickened accordingly. This is one indicator of the presence of monoclonal Ig.
The identification of Ig by means of precipitin reaction is achieved by placing cellulose acetate strips saturated with antisera on the separation lanes. Usually, five antisera are used per patient sample (i.e., anti-Ig/γ-chain, anti-Ig/α-chain, anti-Ig/μ-chain, anti-Ig/L-kappa, and anti-Ig/L-lambda).
After precipitate formation (within 1 h), proteins not precipitated as immune complexes are washed out and the immunoprecipitates are stained using a protein dye. The following patterns can be interpreted:
- Homogeneous band(s) in diffuse immune precipitate: this suggests the presence of a monoclonal or oligoclonal gammopathy
- Diffuse immunoprecipitates: these suggest the presence of a polyclonal gammopathy.
Principle: this method consists of the unidimensional electrophoretic separation of proteins in an antiserum-containing agarose gel. The protein to be detected in the sample is precipitated by the corresponding antibodies in the gel within the equivalence zone in the form of a rocket (rocket electrophoresis). The length of the rocket is proportional to the concentration of the antigen .
Principle: in a first electrophoretic step, the proteins of the sample are separated in agarose gel. By means of a second electrophoresis that is performed at a right angle to the first separation, the single proteins are separated in an antiserum containing gel that contains antibodies to several proteins of interest. Within the equivalence zone, a precipitation occurs with the formation of peaks whose position is determined by the mobility of the protein in the first electrophoretic step and the protein concentration is determined by the height of the peak .
Soluble antigen-antibody complexes absorb and scatter light. The test reagent contains a defined amount of specific antibodies. The antigen concentration in the liquid phase under conditions of antibody excess (i.e., in the ascending part of the Heidelberger and Kendall curve) is measured quantitatively . The limit of detection of this technique, which is around 10 mg/L, can be increased by a factor of 100 if the specific antibodies are bound to latex particles (latex enhanced assays).
Principle: when an antigen containing sample, buffer and specific antiserum are added to a cuvette, antigen-antibody complexes are formed. Light from a helium-neon laser directed through the cuvette is scattered by the immune complexes and the scattered light is focused onto a photo detector by a system of lenses. The electrical signal of the photo detector is proportional to the intensity of the scattered light. The concentration of the antigen can be determined from the scattered light signal by means of a calibration curve. In the case of kinetic nephelometric antigen determination (rate assay), changes in the scattered light are measured at short intervals, while the endpoint method allows the reaction to occur for a defined period of time (e.g., 15 or 30 min.). By adding further antigens or antibodies, it is possible to check whether the measurement occurred on the ascending slope of the Heidelberger and Kendall curve. This is the case if the addition of antigen results in an increase of the measurement signal and the addition of antibody produces no change in the measurement signal.
Principle: when an antigen containing sample, buffer and an excess of specific antiserum are added to a cuvette, soluble immune complexes are formed. In the case of kinetic antigen determination (rate assay), changes in the absorbed light are measured at short intervals. The change in the turbidity of the mixture is measured spectrophotometrically at 334 or 340 nm. An increase in absorption within a defined time period provides a measure of the antigen concentration in the sample (endpoint method). The addition of an accelerator to the mixture accelerates the antigen-antibody reaction so that a kinetic measurement according to the fixed-time principle is also possible.
Indirect antigen or antibody analyses are used if:
- The antigen-antibody reaction does not allow the formation of large immune complexes and, therefore, a visible agglutination cannot be achieved
- A more sensitive detection limit is required than in the aforementioned tests.
Latex agglutination tests can be conducted as slide tests for qualitative determination and turbidimetric or nephelometric tests for quantitative determination of antigens and antibodies.
Principle: the reaction partner of the antibody to be determined in the sample is bound to red blood cells (RBCs) . If the antibody of interest is not present in the sample, no hemagglutination occurs and the RBCs display a dotted or annular pattern of sedimentation. Diffuse agglutination in the form of a mat indicates a positive reaction (presence of the antibody). Quantitative evaluation is possible by titrating the serum sample (dilution with an incubation buffer at fixed ratios).
Direct hemagglutination inhibition test
Principle: certain viruses, in particular rubella viruses, produce agglutinins that agglutinate RBCs (e.g., of chickens) in vitro. If the patient’s serum contains antibodies to viral agglutinins, the agglutination is inhibited.
Indirect (passive) hemagglutination inhibition test
Principle: stabilized antigen-coated RBCs and corresponding antibodies are used as the test system. If the added specimen does not contain the antigen that corresponds to the antibodies, RBC agglutination occurs (negative test result). If the antigen is present in the specimen, it binds to the antibodies and hemagglutination is inhibited (ring or button formation = positive test result).
Principle: antigen-antibody complexes bind and activate complement . The CF test is used for the detection of complement binding antibodies in specimens such as serum or cerebrospinal fluid. The antigen corresponding to the antibody to be measured and a defined quantity of complement are added to the inactivated (complement-free) serum of the patient. If the antibody of interest is present in the specimen, complement binds to the antibody and is thereby totally or partially consumed. Immune complexes (consisting of antibody-coated erythrocytes) that are subsequently added as an indicator system are not hemolyzed, erythrocytes sediment and form a button in the micro titer plate cavity (positive reaction). Complement mediated hemolysis of the indicator system occurs if the specimen does not contain the antibody of interest (negative reaction).
False positive results in the CF test may be caused by antigenic substances (e.g., cell culture antigens during viral cultivation) which may contaminate the true antigen and form a complement fixing immune complex with the antibody of the specimen. Such a false positive result can be recognized by the positive reaction of a control antigen (non infected cell culture material).
More problematic are false positive results due to in vivo immune complex formation (e.g., preformed immune complexes, rheumatoid factors, and aggregated immunoglobulin). This reaction, which is referred to as anticomplementary activity or as serum auto inhibition, is recognized by a positive result in the serum control (patient serum without added antigen).
In infections that trigger a humoral immune response, the proportion of high avidity antibodies increases as the time following immunization increases. This effect is also referred to as maturation of the immune response.
This means that if large quantities of antigen are present in the early stage of an infection, mainly B cells carrying low avidity antibody receptors are stimulated and a population of low avidity poly reactive antibodies is produced.
As time progresses, the antigen concentration declines, which leads to the selection of high avidity antibody producing B-cell clones with additional production of high avidity antibodies. The selection process is closely linked with the production of memory B cells, so that in the case of a reactivated infection or reinfection, high avidity antibodies are produced immediately .
Primary infections have a low proportion of high avidity antibodies and a high proportion of low avidity antibodies. In the first 200 days following primary rubella infection, for example, approximately equal proportions of high avidity and low avidity antibodies are measured. Reactivated infections or reinfection can be distinguished from primary infections based on the proportion of high avidity antibodies (usually > 50%).
Principle of avidity index determination: the stability of antigen-antibody binding is measured in an enzyme immunoassay or the indirect immunofluorescence test. The elution principle is used in the majority of cases. Low and high avidity antibodies in the sample are allowed to bind to an antigen that is bound to a solid phase. After the subsequent washing step, which uses 6 mol/L urea, only the high avidity antibodies remain bound. Two test mixtures with the same sample are analyzed. One test mixture is washed with normal buffer solution. Both high and low affinity antibodies remain bound to the solid phase and are measured. The second test mixture is washed with urea. Only the high avidity antibodies remain bound to the solid phase and are measured. The proportion of high avidity antibodies to total bound antibodies is expressed as avidity index. Depending on the infection, an avidity index > 30% (50–70%) suggests reactivated infection or reinfection.
Immunofluorescence tests are a combination of histological and immunological techniques. These tests are used to detect antigens in tissues, isolated cells, or microorganisms or to detect antibodies in serum directed against antigens of such tissues or cells. A distinction is made between /, /:
- The direct immunofluorescence test; antigens on tissue sections or cells fixed on a slide are assayed using a fluorochrome-labeled specific antibody
- Indirect immunofluorescence test; serum antibodies directed against antigens of tissue cells or pathogens are measured. The serum antibody binds to the cell and the antigen is detected using a fluorochrome-labeled antihuman globulin.
Fluorescence microscopy is used with different light sources and filter options. The fluorescence patterns and intensities are criteria for the presence of a specific antigen or antibody and its concentration.
Direct immunofluorescence test
Principle: for the detection of tissue and cell bound antigens, tissue sections or cell smears are coated with specific fluorochrome labeled antibodies. Commonly used fluorochromes include fluorescein isothiocyanate and rhodamine. Excess conjugated antibody is washed away and the fluorescence of the cell, tissue structure or pathogen is evaluated microscopically.
Indirect immunofluorescence test
Principle: mounted tissue sections or tissue culture cells, fixed on microscope slides, are the source for the antigen. Diluted patient’s serum is added, followed by incubation. Excess serum proteins are washed away. Fluorescein conjugated antihuman polyvalent immunoglobulin (the detecting reagent) is added followed by incubation. Excess unbound conjugate is washed away and the slide is evaluated microscopically for the presence of specific cellular immunofluorescence.
Immunoassay is a type of binding or ligand assay that depends on the antigen-antibody reaction . Ligand-binding assays employ a specific binding protein and the corresponding analyte (ligand). The binding protein can be a receptor, a protein such as vitamin B12-binding protein, or an antibody. If the binding protein is an antibody, the test is referred to as an immunoassay. The immunoassays are divided into two groups according to whether the substance to be tested are antigens or antibodies.
Immunoassays are quantitative methods that are performed in test tubes or in micro titer plate wells. In general, one of the reaction partners is labeled to enable the quantity bound in the antigen-antibody reaction to be determined. Depending on the type of labeling, the tests are classified as radioimmunoassay or non isotopic immunoassay (e.g., enzyme immunoassay, fluorescence immunoassay, luminescence immunoassay) /, /.
The following test principles are distinguished:
- Competitive and noncompetitive immunoassays
- Assays in which the antigen or the antibody is labeled
- Types in which reactions occur in the liquid phase or between solid and liquid phases
- Homogenous and heterogenous immunoassays.
In the classic heterogenous immunoassay the limited amount of antibody used is insufficient to bind all antigen (analyte) of the sample /, /. The marker labeled antigen (tracer) and the analyte in the sample compete for a limited number of antibodies. Because the antibody displays equal avidity for the tracer and the analyte, antibody binding to the tracer is inversely proportional to the concentration of the analyte. Then, the free and antibody bound tracer are separated and the proportion of antibody bound tracer is determined. The proportion of the tracer in the bound form is inversely related to the initial amount of unlabeled antigen (B0). If the tracer is labeled with radioactivity (radioimmunoassay) a dose-response curve is generated for the measurements by determining the ratio of the radioactivity that is measured in the absence of the analyte (B0) to the radioactivity determined in the presence of a known analyte concentration (B) . The higher the ratio B/ B0 the lower the concentration of the analyte in the sample.
Non competitive method
An example of the non competitive method is the two-side sandwich immunometric assay /, /. The antigen of the sample is allowed to react with the solid-phase bound antibodies. In the next step an excess of labeled antibodies which bind to another side of the antigen is added. A separation of bound and free forms is necessary. The portion of the labeled antibodies bound is related to the initial amount of antigen in the test sample.
In the homogeneous immunoassay the extent of antigen-antibody reaction is determined without physical separation of the free antigen and the antibody bound antigen . The term is used for immunoassays such as enzyme and fluorescence immunoassays in which one partner is labeled (e.g., enzyme or fluorescence dye). The assays are designed to measure free antigen as well as antibody bound antigen after the reagents are added in a stepwise manner.
- Immunoassay based on enzyme labeled antigen that competes with the analyte of the patient sample for antibodies contained in the reagents of the assay. Usually the free form of enzyme labeled antigen has higher activity than the antibody bound form. Any change in the activity of the enzyme labeled antigen is related to the concentration of the analyte in the sample. After the addition of substrate, the substrate turnover is proportional to the concentration of the analyte in the sample of the patient.
- Immunoassay based on enzyme modulator or prosthetic group labeled antigen that competes with the analyte of the patient sample for antibodies contained in the reagents of the assay. By binding to an antibody the activity of the enzyme modulator or prosthetic group is inhibited and can be reversed by adding the analyte of a patient sample. Thus the amount of analyte in the patient sample can be determined by measuring the increase in the activity of the enzyme.
- Immunoassay based on reactant labeled antigen: an enzyme and its corresponding substrate-antigen conjugate are used. The reactant labeled antigen competes with the analyte of the patient sample for antibodies contained in the reagents of the assay. The enzyme can bind to the free form of the substrate-antigen conjugate and the substrate is transformed to a product. However, when the antibody is bound to the substrate-antigen (bound form) the enzyme reaction does not take place. When the amount of unlabeled antigen is elevated, the free form of the substrate-antigen conjugate increases and the product of the enzyme reaction is increased.
In the heterogeneous immunoassay, the immune reaction partner is bound to a solid phase (tube wall). A distinction is made between competitive and noncompetitive assays.
Example of a competitive assay
A limited number of antibodies are available, bound to a solid phase. The analyte and a constant quantity of enzyme labeled antigen compete for the solid phase antibody. The higher the concentration of analyte, the less enzyme labeled antigen is bound, and the intensity of the color reaction when the substrate is added is inversely proportional to the analyte concentration in the sample. These assays are referred to as reagent-limited assays.
Example of a non competitive assay
Capture antibodies (in excess) are bound to the surface of a solid phase. Each analyte in the sample binds to these antibodies to form a solid phase bound immune complex. A second labeled antibody is added and a labeled immune complex sandwich is formed. Depending on the procedure used, this type of assay is also referred to as a reagent-excess assay, two-site immunoassay, or two-site (sandwich) immunoassay.
Depending on the number of incubation steps, the following types are distinguished:
- Two-step assay: after the analyte is bound to the capture antibody, unbound sample components are separated and washed away. Then, a labeled second antibody (signal antibody) is added in a second incubation step. The second antibody binds to another epitope of the solid phase bound analyte, forming a sandwich that surrounds the antigen from both sides. The activity of the bound signal antibody is proportional to the analyte concentration in the sample. The two step principle is used in the enzyme linked immunosorbent assay (ELISA).
- One step assay: the reagents are added to the test preparation in a stepwise manner and only one incubation is performed. Capture antibodies and signal antibodies bind to different epitopes on the analyte.
Enzyme-linked immunoassay (ELISA)
Enzyme immunoassays using enzymes as markers are mostly two step assays (). In some cases, the capture antibody loses its binding capacity due to steric hindrance after binding to the solid phase. To prevent this, the solid phase is coated with a species specific antibody, which is then used to immobilize the capture antibody.
Antibody capture assay
In the μ-capture immunoassay, a species specific IgG antibody directed against the Fc fragment of the IgM molecule is bound to a solid phase as a capture antibody (). The concentration of antigen specific IgM antibodies is measured
Enzyme-labeled antigen assay (ELA)
The direct binding of antigen or antibody to a solid phase can result in a loss of sensitivity and specificity of an immunoassay. The streptavidin-biotin method is a system for indirectly fixing antigen or antibody to the solid phase.
A streptavidin coated tube serves as the solid phase. Streptavidin has a capture function and binds biotinylated antibody or biotinylated antigen. Streptavidin is a tetrameric protein with a molecular mass of 60 kDa that is capable of binding four biotinylated antigens or antibodies. Refer to .
The most commonly used radioactive label is 125I, a γ-radiation emitting isotope with a half life of 60 days. The specific radioactivity of the 125I-labeled ligand should amount to a quantity that allows the measurement of several thousand counts per minute (cpm) and per test. One curie (C ) equals 3.7 × 1010 becquerels (Bq); this is equivalent to 3.7 × 1010 disintegrations per second.
Immunoassays with enzyme labels are referred to as enzyme immunoassays (EIA). Either the antigen or the antibody is labeled. EIAs in which the separation technique involves a solid phase are referred to as enzyme linked immunosorbent assay (ELISA). Enzymes that are often used as labels include alkaline phosphatase, horseradish peroxidase, and glucose-6-phosphate dehydrogenase. After the addition of a specific substrate, the enzyme catalyzes the transformation into a product whose formation is determined either kinetically or by means of an endpoint measurement.
Immunoassay with fluorophore labeling is referred to as fluorescence immunoassay (FIA). Fluorophores are molecules that absorb energy and release it again as photons within a time period of about 10–8 sec. If the energy is supplied in the form of radiation, it is released again with a wavelength shift of about 30–50 nm. A commonly used fluorescent dye is 4-methyl umbelliferone phosphate, which is dephosphorylated to the fluorophore 4-methyl umbelliferone by alkaline phosphatase.
The antigen or antibody is labeled with a luminescent substance. The luminescent substance consists of a molecule that emits light after having been supplied with energy.
A distinction is made between the following types of luminescence:
- Chemiluminescence: this is generated by substances that emit light as a result of chemical oxidation, (e.g., luminol, acridinium ester, and oxalate). Catalysts such as horseradish peroxidase or hydrogen peroxide are required to start the chemiluminescence reaction ().
- Bioluminescence: light is produced by a luminescent system that is preceded by an energy providing enzymatic reaction. The luciferin-luciferase system is used to produce luminescence (). Luciferase catalyzes the oxidation of luciferin into an excited state that disintegrates while emitting light. Preceding energy providing reactions include, for example, the ATP providing pyruvate kinase reaction if the luciferin-luciferase system from lightning bugs is used or oxidative reactions if the NAD(P)H dependent luciferin-luciferase system from fungi, algae, or bacteria is used.
With a luminescence immunoassay, substances of low and high molecular weight can be determined with a detection limit that is comparable to that of a radioimmunoassay.
The detection of a labeled substance depends on:
- The number of signals produced within a defined period of time by a defined quantity of labeled molecules (specific activity)
- The ratio of measured signals to signals produced (counting or measurement yield)
- The intensity of the background signal against which the specific signal is measured (signal-noise ratio).
Because of their higher specific activity, luminophores and fluorophores can be detected more sensitively than the radioactive 125I label. A comparison between the detection limits of different immunochemical methods is only possible if important assay factors (e.g., antibody characteristics or separation methods) are kept the same.
Factors or substances that interfere with immunoassays cause differences between the measured and true results. The International Federation of Clinical Chemistry defines interference as follows: analytical interference is the systematic error of measurement caused by a sample component, which does not, by itself, produce a signal in the measuring system .
This interference may be independent of or dependent on the analyte concentration and may increase (positive interference) or reduce (negative interference) the measured result . Positive interference in immunoassays is based on a lack of specificity. The frequency of interference in immunoassays is estimated to be 4% and can be reduced to 0.1% by removing the Fc fragments of the capture antibody .
Circulating human antibodies against animal protein, in particular human anti-mouse antibodies (HAMA) and heterophile antibodies, are the most common causes of interference in immunoassays. Two-site (sandwich) immunoassays are most frequently affected.
The role of heterophile antibodies as interference factors was investigated in a study that performed immunoassays for 74 analytes on serum samples from 10 donors with illnesses known to be associated with rheumatoid factor. Of the 3,445 results 8.5% were considered to be false positive:
- and 21% of these would potentially have led to an incorrect diagnosis
- and 39% would have produced adverse clinical consequences
- and in around half of cases, the erroneously high results were corrected by the addition of blocking reagent.
Interference due to HAMA was investigated in another study in which a two site (sandwich) CEA immunoassay was performed on 11,261 samples. The frequency of interference was estimated to be 4%. The addition of denatured mouse immunoglobulin or the removal of part of the Fc fragment from the capture antibody reduced the frequency of interference to 0.1%.
Despite this apparently high incidence of heterophile antibodies and HAMA, screening of samples prior to immunoassay is not recommended. To date, there is no reliable means of identifying specimens likely to cause interference in immunoassays . Clinical staff must discuss any unexpected immunoassay result with laboratory staff.
- Comparison of current results with previous results
- Discussion with clinical staff and comparison of results with the clinical picture
- Comparison of results with other clinical or laboratory findings relating to organ disease or clinical symptoms (e.g., whether an inverse relationship exists between the aberrant FT4 value and the TSH)
- Comparison with another assay
- Performance of serial dilutions to investigate whether the stoichiometry is correct; a lack of linearity on dilution suggests the presence of interference
- Screening of samples for anti-mouse antibodies
- Pretreatment of samples with blocking antibodies to block anti-animal antibody interference.
Interference can be caused by:
- Pre-analytical conditions. These relate primarily to incorrect samples or incorrect collection of samples. Serum is the specimen of choice. Although lithium-heparin plasma can also be used, its use in many immunoassays has not been sufficiently evaluated.
- Matrix effects; these are defined as the total effect of all qualitative and quantitative components in the system apart from those of the analyte. These components include plasma proteins, heterophile antibodies, rheumatoid factors, and anti-mouse antibodies
- Lack of test specificity. This leads to crossreactivity with substances that have a similar structure to the analyte
- Interference due to other plasma components such as rheumatoid factors (), thyroid hormone autoantibodies (). Plasma components may also interfere non specifically with immunoassays (e.g., digoxin-like substances or a high concentration of free fatty acids in the case of assays for free thyroid hormone).
- Cross-reactive substances ().
- Hook effect; this is a false negative result due to very high antigen concentrations
- Mechanical interference; cryoglobulins and monoclonal immunoglobulins interfere with various steps in the immunoassay, from sample pipetting, through incubation, to the separation of free from antibody-bound labeled antigen.
- Biotin, also named vitamin B7 or vitamin K a water soluble vitamin of vitamin B complex is consumed from some people in doses of 0.3 to 10 mg daily. In immunoassays biotin interferes with streptavidin-biotin. High dosage of exogenous biotin competes with the binding sites of streptavidin. Exogenous biotin causes false low results in sandwich immunoassays. In competitive immunoassays the concentration of the analyte is inverse related to the signal intensity. The concentration of the analyte is incorrectly increased .
Simoa is based upon the isolation of immunocomplexes on paramagnetic beads using ELISA reagents. The main difference between Simoa and conventional immunoassay lies in the ability to trap single molecules in femtoliter-sized wells, allowing for digital readout of each individual bead coupled with enzyme and substrate. When the enzyme label catalyzes substrate conversion to a fluorescent product the resulting fluorophores are confined to the well, creating a measurable fluorescence signal within a short period of time. The field view of the camera encompasses hundreds of thousands microwells; thus, thousands of single-molecule signals in the array can be counted simultaneously The counting of active and inactive wells constitutes a digital signal corresponding to the presence or absence of single protein molecules. Simoa technology can detect proteins in blood at sub-femtomolar concentrations (10–16 moles/L) in comparison to enzyme immunoassays with a limited sensitivity to picomolar range (10–12 moles/L) /, /.
The blotting technique has the following advantages:
- The electrophoretic pattern of the separated proteins is transferred exactly onto the solid matrix in immobilized form
- After transfer, a wide range of analytical procedures can subsequently be applied to the immobilized proteins; in electrophoresis gels, such reactions are difficult or impossible to perform.
From a nomenclatural and historical perspective, three blotting procedures exist:
- Western blotting, also known as protein immunoblotting: this is a procedure for electrophoretic separation and transfer of proteins to a solid phase
- Southern blotting: in this method, electrophoretically fractionated DNA fragments are transferred to a solid phase (usually nitrocellulose) by capillary action
- Northern blotting, a modification of Southern blotting: in this method, small DNA fragments that do not bind sufficiently to nitrocellulose are coupled to diazobenzyloxymethyl paper.
Blotting methods are the basis of fingerprinting. This term describes a sequence of investigations in which a substance such as DNA, is split into fragments, separated, and identified using two dimensional electrophoretic and chromatographic separation. DNA fingerprinting is important in the detection of genetic defects (hereditary diseases), in forensic medicine, in prenatal diagnosis, and for the determination of paternity.
A powerful tool in molecular genetics is the blotting technique of Southern in which electrophoretically fractionated DNA is immobilized onto nitrocellulose strips and used to examine complementary sequences by hybridization in situ. An adaption of the Southern blot is the covalent attachment of fractionated DNA to diazo benzyloxymethyl paper in order to probe for complementary DNA sequences (Northern blotting) .
Protein blotting is the transfer of electrophoretically separated proteins onto immobilizing matrices /, /. The purpose of this transfer is usually to facilitate binding of macromolecular ligands (e.g., antibodies) to the proteins on the matrix. In the large majority of applications the ligand is an antibody and we speak about immunoblotting also referred as Western blotting . Protein blotting, is an immune reaction that takes place in sequential phases (polyacrylamide gel electrophoresis, transfer of antigens, antibody detection on membrane strips containing antigens) and is often used to detect antibodies in body fluids /, /.
A typical immunoblotting experiment may be divided in the following steps):
- Electrophoretic separation of the sample. The main separation techniques used are high resolution two dimensional flat gel electrophoresis or isoelectric focusing. The support media used are agarose or polyacrylamide sheets. The buffers contain sodium dodecyl sulfate (SDS) or urea.
- Transfer of the separated proteins to a membrane. This membrane must immobilize the proteins so that no diffusion occurs. Nitrocellulose strips or diazo benzyloxymethyl paper are used; the latter binds the proteins covalently. The transfer of the proteins from the electrophoresis gel on the membrane (blotting) can take place by means of simple or assisted diffusion (vacuum, forced pressure), by capillary pressure, or electrophoretically (electroblotting). Electroblotting is the most efficient and the most commonly used method. After protein transfer, nonspecific binding sites on the membrane are blocked.
- Incubation with the sample.
- Detection of the bound ligand. The immune complexes thus formed can be detected by protein staining or using an enzyme coupled second antibody (conjugate) and detection of the catalytic reaction by dye formation. The pattern of protein bands is assessed.
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Gregor Rothe, Stefan Barlage, Evelyn Orsу, Gerd Schmitz
Flow cytometry is a method for differentiation and counting of cells in suspension on basis of their fluorescence after binding of labeled antibodies and because of their light scattering properties.
The schematic illustration of a flow cytometer, including an optical unit, a fluid based transport system, and a signal detection unit is shown in . The optical unit consists of a light source, usually laser, a system of lenses for focusing the laser beam, and a fluid cell in which the particles to be analyzed are illuminated by the laser beam. The cells or particles are focused hydrodynamically within a fluid stream and are transported to the point where the laser and liquid stream intersect. The cells or particles become illuminated by the laser beam and emit scattered light. The size of the cell or particle is determined by the forward light scatter and its granularity is determined by the orthogonal light scatter. The fluorescence of cells or particles is also detected at right angles to the incident laser beam.
For selective analysis of a specific fluorescence signal, a system of mirrors and filters is used that enables specific wavelength ranges to be captured on individual detectors.
The photomultiplier tubes (PMT) used to detect fluorescence amplify the signal and convert optical signals into electrical current. The decision about whether to use linear or logarithmic amplification depends on the dynamics of the measurement range, which in the case of immunofluorescence measurements can extend over a number of decades, making logarithmic amplification necessary.
DNA measurements are amplified in a linear fashion. The electrical currents are digitalized by analog-to-digital converters and, depending on the signal intensity, are assigned to a specific channel number (e.g., between 0 and 1023 or 1 and 4096) depending on the resolution. This data is stored as list files (list mode) and can be evaluated for each measured cell in the data analysis.
In the case of single parameter evaluations, the frequency distribution of signal intensities across a cell population is represented in the form of a histogram (). If there are two parameters, these are represented in the form of xy diagrams known as dot plots. Homogeneous cell populations are represented by dot clouds within a two dimensional diagram.
Heterogeneous samples result in complex dot plots with many partially overlapping dot clouds, so special software tools are required to evaluate individual populations. Software featuring largely automated evaluation strategies is available for a range of applications.
Modern flow cytometers are designed to enable the simultaneous use of multiple fluorescence labels, thereby allowing multiparameter analysis. Any overlap between the emission spectra of the fluorescent dyes used can be offset by compensatory measures.
Because monochromatic lasers are employed as the excitation light sources, the number of fluorochromes with corresponding excitation wavelengths that can be excited is limited. For this reason, routinely used devices often feature dual laser excitation.
The expression of cellular antigens can be analyzed by means of immunofluorescence techniques employing fluorochrome labeled antibodies. A selection of diagnostically important antibodies can be found in . These are monoclonal antibodies that have been assigned to antibody clusters (CD, clusters of differentiation) in international workshops based on their specific characteristics . Because of their higher cross reactivity, polyclonal antibody preparations are used in isolated cases only, for example, to detect cell associated immunoglobulin on B cells or bound receptor ligands. Combinations of antibodies can be used to differentiate between leukocytes based on their characteristic antigen expression. Flow cytometry can therefore be used to perform an immunological differential blood count that surpasses that of classical hematology analyzers in its ability to differentiate between T and B lymphocytes as well as natural killer (NK) cells and other T cell populations. Individual cell populations can be defined by analyzing their light scattering properties in combination with data relating to antigen expression or the expression density of individual antigens. A summary of the expression data for selected antigens on cell populations in the peripheral blood is provided in .
Flow cytometry is a valuable addition to existing hematological diagnostic tests:
- By enabling immunologically defined lymphocyte populations to be identified
- By providing a high degree of precision and reproducibility by analyzing large numbers of cells
- By effectively separating and differentiating immature and abnormal cells
- By clarifying unclear findings from morphological or automated WBC differentials as part of a staged diagnostic approach.
Another important diagnostic application of flow cytometry is the evaluation of leukemias, lymphomas, and myelodysplastic diseases /, /. In almost all cases, it is possible to clearly categorize blast cells as being of myeloid or lymphoid cell lineage. When the normal antigen expression pattern in physiological progenitor cells is known, pathological antigen patterns can also be detected, especially if these are characterized by aberrant (co expression of an antigen from a different cell lineage) or asynchronous (expression of an antigen from another stage of differentiation) antigen expression. If malignant cells express a characteristic antigen pattern, this can be used to classify disease and to detect residual cells as part of monitoring.
In addition to the analysis of blood and bone marrow samples, immunophenotyping can also be used to analyze fluids obtained by aspiration (cerebrospinal fluid, ascites, pleural fluid) or lavage methods (bronchoalveolar lavage, BAL) as well as tissue samples. In the case of tissue samples, cell isolation must first be carried out.
Until now, the absolute concentration of individual cell populations has in many cases been determined by calculating their respective percentage proportion of the overall cell population, which was determined using hematology analyzers or other cell analyzers. However, more precise measurements can be obtained directly using a flow cytometer and one of the following approaches:
- Volume calibration (not provided in most commercially available flow cytometers)
- Addition of a defined number of micro particles, based on which the concentration of individual cell populations in the same sample can be calculated.
Analyses aimed at determining the exact concentration of individual cell populations (e.g., the analysis of CD4+T cells or CD34+ progenitor cells) should therefore be conducted using these “single-platform” methods .
In micro particle based test systems, reactions such as antigen-antibody reactions, enzyme-substrate reactions, or nucleic acid hybridization are allowed to take place on the surface of fluorescent particles and quantified. If micro particles with varying fluorescence intensity or different spectral characteristics due to the presence of varying proportions of two dyes are used, the flow cytometer can differentiate between a multitude of different particle populations. If individual particle populations with different coatings are used, multiple analytes can be measured simultaneously in a single sample. Test systems for analyzing specific cytokines and hormones and for use in infection serology are available commercially but their use is often restricted to manufacturer specific flow cytometers .
A single cell suspension is a prerequisite for the analysis of individual cells. Samples of anticoagulated whole blood or bone marrow are therefore required. The anticoagulant used depends on the type of analysis. While EDTA blood samples are frequently used for immunophenotyping, the majority of functional tests require Ca2+-containing media and therefore use heparinized blood samples.
Platelet function tests are conducted using citrated whole blood. Due to the use of multiparameter analysis, enrichment of cell populations (e.g., by density gradient centrifugation) is not performed in most cases since the relevant populations can be detected using specific labels . Cell containing aspirates (cerebrospinal fluid) and lavage fluid (BAL) can also be analyzed. Tissue samples can be analyzed, following isolation of the relevant cells.
Blood samples are transported and stored at room temperature in most cases. However, samples with reduced cell stability, such as cerebrospinal fluid or other aspirates, should be transported and stored at a low temperature. For functional tests, samples must be transported to the laboratory, prepared, and analyzed as quickly as possible.
Samples for immunophenotyping should be processed on the same day; prolonged storage or transport of samples by mail can limit the significance of test results for the respective analyses.
Methods for stabilizing cells without fixation open up new possibilities by allowing even complex immunophenotyping to be performed on samples that are up to a week old.
Body position during blood collection
For the quantitative determination of cell populations, it is important to note that the concentration of corpuscular blood components is 5–10% lower if the blood sample is collected when the patient is in supine position.
Time of day
Many values exhibit a circadian rhythm.
The distribution of lymphocyte populations in the peripheral blood is age dependent.
Depending on the test in question, many other factors such as lifestyle habits (e.g., nicotine abuse in the case of bronchoalveolar lavage) or drugs (e.g., masking of epitopes by therapeutic antibodies) may need to be considered.
Problems during cell immunophenotyping
The problems can be due to a wide range of factors:
- Antibody concentrations that are too high can lead to an increase in nonspecific color reactions while concentrations that are too low can prevent quantitative labeling of the epitopes present. Antibody titration using cells with maximum antigen expression is therefore recommended.
- Washing steps, particularly following cell fixation or if low protein buffers are used, can lead to selective cell loss. The use of less adherent test tubes, methods for erythrocyte lysis without fixation, and the addition of protein or calcium chelators to the washing solution can reduce cell loss in some cases.
- Interference due to high cellular auto fluorescence can be reduced by using dyes with an emission spectrum of a longer wavelength or by quenching the auto fluorescence with crystal violet
- Nonspecific binding of antibodies to Fc receptors can be limited by pre incubating cells with excess nonspecific immunoglobulins or incubating them in serum or whole blood
- Stained samples carried over to subsequent samples may mimic abnormal cell populations. This can be recognized by comparing the “dot plots” with those of the previous sample.
- Micro particles used to directly determine absolute cell counts can aggregate at low protein concentrations, leading to incorrect count results. For this reason, buffer solutions with a sufficiently high protein content are required if dilute samples (e.g., from apheresis products) are used.
As for other laboratory methods, internal and external quality assessments should be carried out where available. Ideally, both device checks and process checks to detect errors in sample processing should be carried out. In addition to the fluorescent micro particles used to check device settings, cellular control materials are also available for some applications, which allow sample processing for the analysis of CD4+T lymphocytes to be checked. Inter laboratory surveys are offered for a range of applications such as the determination of lymphocyte sub populations, leukemia and lymphoma diagnosis, reticulocyte analysis, and analysis of CD34 progenitor cells.
Internal plausibility checks are possible for multiparameter measurements, provided the antigen expression patterns of the cell populations in the sample are known. In the analysis of blood samples, for example, the number of cells (B lymphocytes) expressing CD19 and the number expressing CD20 should be approximately equal. The total number of CD4+CD3+T cells and CD8+CD3+ T cells should correspond to the number of CD3-positive T cells. The proportions of T, B, and NK cells should add up to 100% (lymphocyte population).
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Amplification techniques developed for the detection of nucleic acids form the basis of a wide range of diagnostic methods. The polymerase chain reaction (PCR) is currently the most widely used amplification technique. However, the alternative technologies described here are also employed in commercial test kits. Based on the approach used, a distinction is made between amplification of the target nucleic acid sequence (e.g., PCR) and signal amplification without prior DNA amplification (e.g., branched DNA technology). Methodological developments in amplification technology are advancing rapidly, with an increasing emphasis on the quantitative determination of target DNA or RNA sequences and on the use of closed tube formats to prevent contamination.
The PCR has revolutionized nucleic acid based diagnostics within the space of a few years. Thanks to PCR, it was possible to solve one of the most pressing problems in genetics (i.e., how to amplify tiny quantities of DNA). PCR was developed by Kary B. Mullis in 1983 and first published in 1985 . Mullis’s basic idea was to copy DNA from oligonucleotides (primers) in a number of replication cycles using the enzyme DNA polymerase.
The original technique was highly inefficient and susceptible to interference. The DNA polymerase, which has an optimum temperature of 37 °C, was destroyed by the high heat (approximately 95 °C) used at the start of each cycle to denature the DNA. As soon as scientists started to use DNA polymerases isolated from thermophilic bacteria that live in hot springs, the amplification process became significantly easier. Thermus aquaticus is the most prominent microbial source of thermostable DNA polymerase, which consequently was given the name Taq polymerase . In the years that followed, the use of PCR spread throughout the world and ushered in the age of genomics. It should be added that in 1991, just 6 years after the method was published, the Roche company acquired an exclusive license for the method from Cetus (Mullis’s employer at the time) for 300 million dollars, a gigantic sum at the time. As a consequence, commercial use of the technology was initially protected by a worldwide patent. This patent expired in 2005 in the USA and 2006 in Europe, so standard PCR technology is now available for general use. However, the patent extends to numerous follow-on applications, restricting their use. The inventor of PCR was awarded a number of important scientific accolades in record time, culminating in the Nobel Prize in Chemistry in 1993.
PCR is used to amplify short, precisely defined sections of a specific DNA fragment. These fragments are often just a few hundred base pairs (bp) in length; however, fragments of up to 40 kbp (40,000 bp) can also be replicated using certain techniques.
- DNA template that contains the section to be amplified
- Two primers that mark the beginning and end of the section to be amplified
- DNA polymerase, which synthesizes a new DNA strand complementary to the DNA template by adding nucleotides. The DNA polymerase used is thermostable Taq polymerase. The Taq polymerase adds a non complementary nucleotide every 500 bp or so during the amplification process. In the standard procedure, this is usually irrelevant. If, however, these mismatches are a problem, thermostable polymerases with a correction mechanism (e.g., Pwo or Pfu) should be used.
- Nucleotides, which serve as the building blocks for synthesizing the complementary DNA strand
- Buffer solution to ensure suitable reaction conditions.
The standard PCR consists of a series of around 30 recurring cycles, each consisting of three steps: denaturation, annealing, and elongation (). PCR takes place in thermal cyclers, which quickly and precisely regulate the temperature required for each step. Theoretically, the DNA is doubled in each cycle; therefore, the quantity of DNA increases exponentially. In 30 cycles, for example, a billion (230) copies should be produced. The actual quantity is significantly lower, however, because the PCR becomes less efficient with each cycle.
When double stranded DNA is heated to around 95 °C, the hydrogen bonds that hold the complementary DNA strands together are reversibly broken. In the first cycle, the DNA is commonly heated for 3 min. at 95 °C; in subsequent cycles, 30 sec. is sufficient.
As the temperature is subsequently reduced, the primers bind to their complementary sequences on the DNA template or amplified fragments. The annealing temperature of the primers depends on their base sequence and their length. A temperature that is 5 °C below the melting point of the primer is normally chosen as the annealing temperature. Because primers with a length of 20 bp are commonly used, the annealing temperature is usually around 55 °C. However, the optimum annealing temperature for each PCR must be determined experimentally.
The temperature is increased to 72 °C, which is the optimum temperature for thermostable DNA polymerase. DNA polymerase synthesizes complementary DNA strands, starting from the primers.
DNA template: in theory, only one DNA template molecule is required for amplification. However, for consistently successful PCR under routine conditions, at least 10,000 molecules are required. This corresponds to approximately 30 ng of human genomic DNA. The purity of the DNA should also be checked (optical density 1.8 to 2.2).
PCR buffer: the buffer is usually provided by the polymerase manufacturer. Buffer optimization is therefore unnecessary in all applications.
MgCl2: Mg2+ affects primer annealing, the separation of DNA strands during denaturation, product specificity, the production of primer dimers, and the error rate. The polymerase also requires Mg2+ to express its activity. Many components of PCR contain EDTA, which binds the Mg2+ added. A standard concentration of 2 mmol/L is used in PCR applications. To increase the yield or specificity, it may be necessary in individual cases to optimize the Mg2+ concentration. In general, the higher the Mg2+ concentration, the more intensive the PCR product, with a simultaneous decrease in specificity.
Additives: they may be required to increase the specificity, especially for the amplification of GC rich sequences. Commonly used additives include dimethyl sulfoxide (DMSO) (5 to 10%), betaine (N,N,N-trimethylglycine) (1 mol/L), and formamide (up to 5% v/v). Glycerin (10–15% v/v), PEG 6000 (5–15% w/v), and Tween®20 (0.1–2.5% v/v) are used to accelerate the reaction.
Nucleotides: usually, a standard concentration of 200 μmol/L of each nucleotide is used. However, it is important to note that the quality may vary between different manufacturers; testing may therefore still be required.
Annealing temperature: the annealing temperature of the primer is crucial to the yield and specificity. Nowadays, different formulas are used to calculate the melting temperature . The melting temperature is the temperature at which 50% of the primer is no longer bound to the template. The annealing temperature must therefore be 5 to 10 °C lower than this. Despite the relevant calculations, practical testing is often unavoidable. This testing can be performed easily using commercially available gradient PCR devices.
Thanks to the use of commercial systems, PCR optimization has become significantly easier. Some companies offer a whole panel of different kits for PCR applications. In addition to standard PCR kits, there are kits that are optimized for PCR fragments of up to 40 kbp and multiplex PCR. In the case of recurrent nonspecific bands, a HotStar polymerase can be used. The Taq polymerase is activated in an initial activation step at 95 °C. This prevents nonspecific primer annealing and the formation of primer dimers.
RNA cannot be amplified directly using PCR; it must first be transcribed into DNA. In vivo, DNA is normally transcribed into mRNA. The reverse process (the transcription of mRNA into DNA) is therefore referred to as reverse transcription. Reverse transcription is primarily used in the diagnosis of RNA viruses, such as human immunodeficiency virus (HIV) and the hepatitis C virus (HCV), and in the analysis of gene expression. Reverse transcription takes place upstream of PCR by means of reverse transcriptases (RTases). Three reverse transcriptases are available, which differ in terms of their optimal temperature and the length of the RNA segment transcribed :
- AMV (avian myeloblastosis virus) RTase, optimal temperature 42–60 °C, fragment length < 6000 bases
- MMLV (Moloney murine leukemia virus) RTase, optimal temperature 37 °C, fragment length < 20,000 bases
- Tth DNA polymerase, optimal temperature 68–80 °C, fragment length < 1,000 bases.
The most suitable RTase is selected based on the application in question. The reverse transcription step extends a primer complementary to the target RNA sequence or, frequently in the case of mRNA, a universal oligo (dT) primer with a length of 15–20 bases. An important practical distinction exists between one-step and two-step reverse transcription.
In the two-step reaction, reverse transcription and amplification take place separately. This may include using different buffer systems that are optimized for the specific enzyme used.
In the one-step reaction, although reverse transcription takes place before the amplification step, both reactions occur in the same tube. For molecular diagnostics, the one-step reaction is the preferred approach: the lack of a second step simplifies the procedure and the reaction tube does not need to be opened, which reduces the risk of contamination. For scientific applications, however, the two-step approach is used. A particular advantage in this context is that multiple different amplifications can be performed on the DNA generated in the first step by reverse transcription.
Nested PCR approaches increase both the detection limit and specificity of PCR. It comprises two sequential PCR reactions. An aliquot of the first amplicon is used as the template for the second PCR reaction. The primers of the second PCR hybridize to the first amplicon, resulting in a shorter fragment. Due to its high detection limit, nested PCR is associated with an increased risk of contamination. Special precautionary measures and controls must therefore be implemented.
Methyl cytosine is referred to as the “fifth base” in the human genome. It cannot be identified using standard PCR and sequencing. The identification and quantification of methyl cytosine has become the focus of attention in recent times due to the association between methylation and promoter activity in individual genes. Methylation based biomarkers are now available for cancer screening. A blood based assay that detects tumor DNA hyper methylation in the septin 9 gene is commercially available as a screening test for colorectal carcinoma .
The pretreatment of DNA with bisulfite frequently forms the basis of genetic tests. Treatment of single stranded DNA with bisulfite leads to deamination of unmethylated cytosine residues to the RNA base uracil. In the subsequent PCR, uracil is amplified as thymidine. Unlike unmethylated cytosine, methyl cytosine is not converted to uracil. The degree of methylation at a given position can be derived from the ratio of cytosine to thymidine.
shows the pyrosequencing method for analyzing DNA methylation. The nucleotides are pipetted into the reaction mixture according to the sequence of the template. By adding cytosine and thymidine at the position of interest, the degree of methylation can be derived.
Multiplex PCR is a modification of PCR in which several gene loci are amplified simultaneously in one reaction. It provides a rapid and sensitive method for use in research and diagnosis. Compared to the amplification of a single gene locus, the design of a multiplex PCR assay requires significantly more resources. It is essential to ensure that all loci are amplified equally efficiently. Several commercial test systems are available for the molecular diagnosis of sepsis . These assays can detect more than 20 different pathogens that cause more than 90% of all sepsis cases.
Qualitative PCR usually follows a standard protocol. In many cases, specific genes are copied in order to detect disease related changes. PCR was used for the first time to diagnose a disease (sickle cell anemia) in the mid-1980s. At around the same time, the method was introduced to forensic medicine as a means of identifying individuals. Qualitative PCR is also used to detect pathogens. Due to its high limit of detection, it can detect infection at an earlier stage than other methods such as cultures or antibodies.
PCR products are often characterized by size. The most widespread application of PCR is agarose gel electrophoresis, which is both easy to use and cost effective. DNA can be visualized in the agarose gel using ethidium bromide a dye that intercalates between DNA base pairs and fluoresces under ultraviolet light . If a greater degree of resolution of the PCR product is required, polyacrylamide gels or capillary electrophoresis are used.
Following PCR, restriction enzymes are often used to detect mutations. Restriction enzymes cut DNA at or near specific sequences of 4–6 nucleotide bases known as restriction sites. Point mutations can result in an increase or decrease in the number of restriction sites. The presence of mutations can be inferred based on the pattern of fragments in the gel ().
As the use of molecular diagnostics has increased, the methods employed have been simplified. Depending on the manufacturer of the devices and reagents for a particular test, this is achieved in different ways. A common feature of genetic tests, however, is a homogeneous design. PCR and subsequent detection take place in the same tube. The time consuming detection of PCR fragments (e.g., using agarose gel electrophoresis) is no longer necessary. The risk of contamination is avoided because amplification and detection are performed in a closed tube.
In addition to the mere detection of fragments, the use of mutation specific probes also enables allelic differentiation. Two different methods with corresponding analyzers have been widely adopted. In the first approach, two primers and two probes are used in the PCR. The two outer primers are responsible for amplification while the fluorescently labeled inner probes are used to display the PCR fragment and to distinguish between alleles.
This method is based on FRET (fluorescence resonance energy transfer) technology. If the two fluorescent dyes used to label the 5’ end of one probe and the 3’ end of the other are within a certain distance of each other, the excitation energy of the first fluorophore is transferred to the second fluorophore. Measurement is based on light of a specific wavelength that is emitted by the second fluorophore. The inner probes are complementary to the wild type allele, so the melting point is relatively high. If a mutation is present, the fluorescently labeled probe is no longer completely complementary and the melting point decreases accordingly. Melting point analysis is used to distinguish between alleles () .
The second method uses the TaqManTM system. Two primers are used for PCR and a probe located in the fragment is used for detection. Typically, a fluorophore is attached to the 5’ end of the probe and a quencher molecule is attached to the 3’ end. Quencher molecules absorb fluorescence from dyes located in their vicinity. Through the 5’ → 3’ exonuclease activity of Taq polymerase, the fluorophore is separated from the quencher of the probe and upon excitation can emit light of the expected wavelength. To detect nucleotide replacements, a different fluorophore is attached to two probes: one that is typical for the wild type allele and one that is complementary to the mutation. The exonuclease activity of Taq polymerase is only active if there is 100% complementarity between fragment and probe. The probe that is complementary to the wild type allele emits light of a particular wavelength only if the wild type allele is present. In the case of heterozygosity, the fluorescence signal is correspondingly reduced by half. If only the mutation is present, no fluorescence signal is emitted. The probe for the mutation behaves in the same way: it emits a fluorescence signal only if the mutation is present .
Quantitative PCR detects whether a particular DNA sequence is present in a sample as well as how much is present . Quantitative PCR is mainly used in medical diagnostics and research. It is based on the fact that the amount of reaction product is doubled in each cycle, which allows the initial quantity to be inferred. The actual efficiency of PCR lies below this theoretical value, however. This is because the replication rate is not always optimal, particularly in the first and last PCR cycles. This means that the quantity of DNA molecules initially present cannot be inferred based on the quantity of DNA at the end of PCR.
Many different approaches to quantitative PCR have been described. Real-time PCR is considered to be the gold standard in quantitative PCR . The devices used for real-time PCR employ optic fluorescence and can therefore measure the quantity of DNA produced in each cycle. The amplified DNA can be detected in two ways: using specific fluorescently labeled probes or using fluorescent dyes such as SYBR Green that react with any double-stranded DNA. Fluorescently labeled probes detect only the sequence of interest whereas intercalating DNA dyes detect any double-stranded DNA, including any nonspecific amplification products. In quantitative PCR, therefore, fluorescently labeled probes that detect only the DNA sequence of interest are preferred.
The PCR product can be quantified in absolute or relative terms. If absolute quantification is used, a sample with a known number of target molecules must be used as a reference. In the case of relative quantification of the target sequence, a stable reference gene (housekeeping gene) is used. Quantitative PCR is becoming increasingly important in the diagnosis of infectious diseases (). Relative quantification is primarily used to analyze cellular gene expression patterns.
The high detection limit of PCR (in theory, only one DNA molecule is required) also leads to a risk of contamination with amplified DNA segments, which can lead to false positive results. The prevention of contamination is one of the main challenges faced by genetic laboratories. Pre-PCR and post-PCR work areas should be physically separate. Sample and reagent preparation take place in the pre-PCR room while amplification and detection of PCR products take place in the post-PCR room. As mentioned above, the use of real-time PCR devices reduces the risk of contamination because PCR and detection take place in the same closed tube. In addition, uracil-N-glycosylase (UNG) is commonly used to destroy previously amplified gene fragments . For this, dTTP in the reaction mixture must first be replaced by dUTP. The reaction mixture is incubated with uracil-N-glycosylase prior to the PCR step. All DNA strands that contain dUTP are degraded. Because the DNA template does not contain dUTP, it remains unchanged; only contaminants from previous amplifications are destroyed. The heat-labile uracil-N-glycosylase is completely inactivated in the first denaturation step at 95 °C.
Detection of hereditary diseases
More than 10,000 disease are known to have a genetic basis . Genetic tests are available for around 600 different hereditary diseases. Although entire chromosomes or segments may be involved, hereditary diseases are most commonly caused by mutations in single nucleotides. If the nucleotide replacements responsible for the disease have been characterized in the gene, the corresponding gene segment can be specifically amplified using PCR and the mutation detected using suitable methods. Because genetic tests are so easy to perform, the range of tests available has expanded considerably. In the case of monogenic disorders with high penetration, the link between the genetic findings and the disease is easy to demonstrate; the same is not true for more complex disorders.
Pharmacogenetics is concerned with inherited genetic differences that influence the pharmacokinetic and pharmacodynamic characteristics of drugs and can affect individual responses to drugs. Many of these genetic differences are phenotypically relevant nucleotide replacements .
If the frequency of this type of variation is more than 1%, it is considered to be a SNP (single nucleotide polymorphism). PCR forms the basis of SNP diagnostics. Of particular interest at present in the area of molecular diagnostics are variations in the cytochrome P450 enzymes CYP2C9, CYP2C19, and CYP2D6. Clinically relevant genetic variations occur in each of these enzymes. A panel containing a range of different mutations is therefore an essential component of pharmacogenetic testing. A commercial CYP450 DNA chip test can simultaneously test for 31 genetic variations in the enzymes CYP2C19 and CYP2D6.
Genetic testing in oncology has several objectives. Although the main focus to date has been on hereditary diseases that predispose affected individuals to the development of cancer, prognostic and therapeutic applications are becoming increasingly important.
Mutations in the KRAS gene are associated with a poor prognosis in colorectal carcinoma and non small cell lung cancer. Patients also show a poor response rate to anti-EGFR therapy (treatment of epithelial carcinoma using anti-EGFR antibodies). It is important to efficiently analyze as many as possible of the multitude of genetic variations that influence prognosis and response to therapy and which may be located on different genes. For this reason, DNA chips are increasingly being used to simultaneously detect a large number of genetic changes.
A relatively new diagnostic area in oncology is the analysis of promoter methylation . In vivo methylation involves the attachment of a methyl group to cytosine. Promoter methylation can suppress the expression of a gene.
The deactivation of tumor suppressor genes is particularly important in oncology. Bisulfite treatment of DNA converts non methylated cytosine into uracil. PCR can then be used to amplify the corresponding promoter segments of the suppressor genes and quantify the degree of methylation.
Forensic DNA analysis
The main application of forensic DNA analysis is genetic fingerprinting, which is primarily used to identify individuals and establish biological relationships. Until the mid-1990s, the main focus was on analyzing polymorphic red cell membrane systems, protein systems, enzyme systems, and the HLA system. The use of DNA technology in association with PCR, however, has revolutionized this area of application .
Forensic DNA analysis is based on the repetition of DNA sequences of variable length and number. The length and number of these repeated segments, also known as short tandem repeats (STR) or micro satellites, vary considerably from person to person.
The short tandem repeats used most commonly in forensic genetics are tetra nucleotide repeats with an average of 10–20 repeating sequences. The length of the PCR products is then determined electrophoretically in high resolution polyacrylamide gel or often by capillary electrophoresis in automated DNA sequencers.
Forensic genetic analyses are subject to strict requirements and regulations. For paternity testing, the German Medical Association stipulates that a minimum of twelve STR markers from at least ten different chromosomes must be amplified. Molecular genetic trace analysis is based on the eight STR marker systems specified in the DNA analysis database of the Bundeskriminalamt (Federal Criminal Police Office). Since 1998, the DNA profiles of crime scene traces, criminal suspects, and convicted criminals in Germany have been analyzed using these 8 marker systems and stored in a database. The intensive use of genetic methods in forensic medicine is illustrated by the high number of data records relating to individuals (570,000) and crime scene traces (142,000) that had already been created by June 2008.
Molecular diagnostic methods are becoming increasingly important in the area of viral and bacterial diseases (refer to and ). If it is only necessary to detect the presence of a pathogen, qualitative PCR is sufficient. For example, mandatory testing for blood donors has been in place for the hepatitis C virus since 1999 and for human immunodeficiency virus type 1 (HIV-1) since 2004.
If the pathogen count is also relevant, quantitative PCR methods are used. The pathogen count is of particular interest in chronic viral infections . It provides more information about disease progression than the clinical features or biochemical parameters.
Quantitative PCR has an important role in assessing therapeutic success, for example, in the case of hepatitis C. The viral load can be used to determine whether the disease is progressing or responding to therapy (refer to ).
Quantitative PCR also has an important role in acquired immunodeficiency syndrome (AIDS), particularly in the area of treatment monitoring. An increase in the viral load may be the first sign that treatment is no longer effective (refer to ).
PCR may also form the basis of a preemptive treatment strategy. In immunosuppressed patients, Cytomegalovirus infection may be a life threatening complication. For this reason, viral load monitoring is commenced in asymptomatic patients and therapy is initiated immediately as soon as the viral load increases.
Branched DNA technology
In this method, the target DNA or RNA is quantified directly without a previous PCR amplification step . It employs signal amplification rather than target amplification. The target DNA or RNA is immobilized by binding to a set of oligonucleotide target probes bound to a solid phase. A second set of oligonucleotides binds to the target sequence captured on the solid phase. These oligonucleotides serve as binding sites for signal amplification molecules. Signal amplification molecules are synthetically constructed DNA molecules with 15 branches, each of which contains three binding sites for alkaline phosphatase-conjugated detection oligonucleotides (). One branch can therefore bind up to 45 detection oligonucleotides. Signal detection takes place in a luminometer by means of chemiluminescence.
Branched DNA technology is used primarily in the diagnosis of viral infections due to HCV and HIV-1. Unlike PCR amplification techniques, branched DNA assays do not require reverse transcription. The oligonucleotides in the reaction mixture can bind directly to the corresponding RNA target sequences of both viruses. The measurement signal is generated by a chemical reaction alone without the need for an enzyme-mediated step.
The viral load is determined quantitatively using a calibration curve. Because they do not involve enzymatic reactions, branched DNA assays are less vulnerable to inhibitors in the reaction mixture. By selecting the oligonucleotides that bind to the target sequence accordingly, it is also possible to capture virus subtypes. Because only the signal is amplified, the contamination risk is significantly lower than in methods that use target sequence amplification.
The advantages of branched DNA technology must be weighed against a slightly lower detection limit in comparison to methods involving gene amplification. In the USA in particular, branched DNA technology is widely used in the diagnosis of HCV and HIV-1; in Europe, however, PCR-based tests are the preferred approach. DNA branched technology in combination with multi analyte profiling beads technology is offered in a variety of kits for simultaneously quantifying different RNA targets in the research context .
Hybrid capture technology
Hybrid capture technology is a method for directly detecting target DNA without a previous enzymatic reaction . It offers the same advantages as the branched DNA method: ease and speed of use. No time consuming DNA preparation is necessary; just a simple chemical reaction to release the DNA. Because target sequence amplification does not take place, hybrid capture technology is also associated with a significantly lower risk of contamination. Released target DNA combines with RNA probes to form DNA-RNA hybrids. The inside of the incubation vessel is coated with antibodies against DNA-RNA hybrids. The DNA-RNA hybrids bind to these antibodies and are immobilized on the wall of the tube. In the next step, an antibody conjugated to alkaline phosphatase binds to the immobilized DNA-RNA hybrids. The signal resulting from the binding of large numbers of enzyme-labeled antibodies to the hybrids is amplified 3000-fold (). The actual measurement signal is generated by a chemiluminescence reaction.
Hybrid capture technology has established itself primarily in the diagnosis of Human Papillomavirus in cervical cancer screening. While PCR is a significantly more sensitive detection method, this degree of sensitivity is not required in this context. Instead, hybrid capture technology is used in association with a clinically relevant cutoff value.
Nucleic acid sequence-based amplification (NASBA)
NASBA typically involves the action of three enzymes . Unlike in PCR, the amplification reaction takes place at a constant temperature of 41 °C and uses single stranded RNA as the starting material. The method is therefore particularly suited to the detection of RNA viruses or pathogen specific mRNA because the reverse transcriptase step that is required at the start of PCR can be omitted. Another advantage of being able to omit the initial reverse transcription step is that RNA amplification takes place against a background of double-stranded DNA.
In the first step of the NASBA reaction, a primer attaches to the target RNA using the complementary binding region (). This primer also contains the T7 polymerase promoter sequence on its 5’ end. Starting from the bound primer, reverse transcriptase synthesizes a complementary cDNA.
In the second step, RNase H selectively degrades the RNA template, leaving behind single-stranded cDNA.
In the third step, a second complementary primer then binds to the single DNA strand and reverse transcriptase synthesizes a second DNA strand. The resulting double-stranded DNA contains the T7 polymerase promoter sequence.
The three reaction steps that lead to the synthesis of the double stranded DNA constitute the initial non cyclic phase. During the cyclic phase, complementary RNA is synthesized by T7 polymerase, starting from the T7 polymerase promoter. Several thousand RNA molecules can be synthesized from a single DNA molecule. The primer binds to the RNA, DNA is synthesized by reverse transcriptase, the RNA strand is degraded by RNase H, the primer binds to the T7 promoter sequence, and double stranded DNA with the corresponding promoter activity is formed. The double stranded DNA then serves as the starter material for the cyclic phase (second strand synthesis). The high amplification efficiency of this method is due to the combination of the non cyclic phase, cyclic phase, and second-strand synthesis.
Transcription mediated nucleic acid amplification (TMA)
TMA is another transcription based amplification method that is similar to NASBA. However, TMA uses only two enzymes. RNase can be omitted because reverse transcriptase has sufficient ribonuclease activity. The use of molecular beacons has significantly simplified and improved the quantification of RNA strands. When molecular beacons are used, fluorescence signals are recorded continuously, which allows quantitative measurement . Molecular beacons are hairpin shaped DNA structures that consist of a stem portion that is complementary to itself and a loop region that contains sequences that are complementary to the target sequence. A chromophore is attached to each end: a fluorophore (usually fluorescein) that acts as a donor and an acceptor (usually dabcyl) that absorbs fluorescence. If the fluorophore and acceptor are in close proximity, the acceptor quenches the fluorescent emission of the fluorophore. However, when the molecular beacon binds to its target sequence, the hairpin structure opens out. The chromophores are separated from each other and the acceptor no longer quenches the fluorescence emitted by the fluorophore. The intensity of the fluorescence signal measured is proportional to the quantity of the target RNA sequence.
Commercially available assays based on NASBA or TMA technology are primarily used in the diagnosis of Human Papillomavirus. Compared to DNA-based assays, these RNA-based assays may be able to discriminate more effectively between transient infection and clinically relevant disease.
In the ligase chain reaction (LCR), two adjacent oligonucleotide probes bind to the target sequence. A thermostable ligase catalyzes the formation of a bond between the 5’ phosphate and 3’ hydroxyl termini of the two probes. From the two oligonucleotides, a new oligonucleotide is formed whose length corresponds to the sum of the lengths of the two original oligonucleotides. Following a denaturation step, the resulting short oligonucleotides bind again to their complementary sequences and the ligase chain reaction cycle repeats itself . The use of different labels on the free outer ends of the ligated oligonucleotide enables the immobilization of the product on the solid phase and the initiation of the detection reaction by signal identification. A disadvantage of the ligase chain reaction described is its lack of specificity compared to other methods. The specificity is often improved by positioning the oligonucleotides a few base pairs apart and using a fill-in reaction to close the resulting gap. Although LCR technology still has a minor role in assays developed in-house, it has been largely superseded in commercial assays.
The four non PCR based amplification methods described often require specialist technology that is not available to ordinary users. For this reason, they are used mainly in commercial applications. Once optimized, these commercial assays are easy to use. However, the number of commercially available kits is decreasing constantly in favor of PCR based methods.
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Mass spectrometry has revolutionized analysis of the human proteosome and improved the clinical assays throughput and precision. However, no HPLC-MS based device for measuring proteins and peptides in in-vitro diagnostic devices has yet been cleared or approved by the Food and Drug Administration (FDA). Proposals are made how further communications and collaboration with the clinical mass spectrometry communities can identify opportunities. An overview of the FDA’s experience regulating MS based devices is published .
Chromatographic methods are used to separate the individual components of a mixture based on their differential adsorption at defined surfaces. In a highly specific separation column, the components are eluted in a characteristic temporal sequence depending on the strength of the interaction between the analyte, the mobile phase, and the stationary phase. The chromatogram represents the results of chromatography, with time on the x-axis and signal intensity on the y-axis (). A cleanly separated substance produces a peak. Ideally, this peak should be symmetrical and should exhibit a constant retention time between samples that enables the substance to be identified. To quantify a substance, the area under the peak or the height of the peak is measured and related to the corresponding value for a calibration sample. In principle, multiple different analytes can be quantified in a single chromatographic run (e.g., epinephrine, norepinephrine, and dopamine in the diagnosis of pheochromocytoma). Chromatography is usually preceded by analyte specific sample preparation in which interfering matrix components are removed and target analytes are enriched and sometimes chemically transformed (derivatization).
Of the multitude of different techniques available in chromatography and mass spectrometry, gas chromatography with mass spectrometric detection (GC-MS), high performance liquid chromatography (HPLC) with various detection methods, and liquid chromatography-tandem mass spectrometry (LC-MS-MS) are most commonly used in laboratory medicine.
Gas chromatography can be used to analyze substances that are volatile or amenable to chemical derivatization to render them volatile. Purified, mainly derivatized samples are vaporized at temperatures of more than 70 °C in the injector of a gas chromatograph and carried to a separation column in a stream of helium (approximately 1 mL/min.; mobile phase). The separation column consists of a coiled flexible quartz capillary of around 30 m in length whose inner surface is coated with a selected substance (stationary phase) and whose inner diameter is around 0.3 mm. During the separation run, which normally lasts around 20 minutes, the temperature of the column oven is increased (e.g., from 70 to 280 °C); temperature gradients can be used to vary the separation characteristics.
The eluate from the gas chromatography is transferred directly to the ion source, which is located in an ultra high vacuum. Here, electrons are released from a hot filament and accelerated toward a cathode. This causes them to collide with the gaseous sample molecules. As a result, the substance disintegrates into fragment ions with different charges (electron impact ionization, EI). The mass spectrometer is configured as a quadrupole consisting of four metal rods arranged in parallel or an ion trap consisting of an trap space surrounded by a ring electrode.
The mass spectrometer acts as a mass filter by applying specific radio frequencies that allow only fragments with a certain mass-to-charge ratio (m/z) to reach an ion detector at a given time while all other ions are deflected. The ions filtered at a specific time strike an aluminum block in the detector to produce electrons. These are amplified in a photomultiplier and the resulting current is measured.
Because the MS radio frequencies can be fine tuned within fractions of a second, it is possible to scan a wide mass range in this period (10–800). As a result of electron impact ionization, most molecules tend to disintegrate into one of 3–5 thermodynamically favored typical fragment ions. The mass spectrum of a substance plots the relative intensity of each of these fragments on the y-axis against the mass-to-charge ratio on the x-axis (). Because mass spectra exhibit relatively high substance specificity and can be compared with extensive spectral libraries, they can be used to identify substances that elute in an individual gas chromatography peak. From the data in the individual mass spectra of an analysis run, the signal of an individual fragment ion mass that is characteristic for a substance can be extracted over time as an ion chromatogram and used to quantify the substance (selected ion monitoring, SIM).
By combining very high separation efficiency with mass fragmentography, GC-MS provides a particularly high degree of analytical specificity. For this reason, it has come to play a key role in toxicology and environmental medicine. The stable isotope dilution technique is the approach most commonly used in quantitative GC-MS. In this technique, an identical quantity of the target analyte, in which several atoms have been replaced by stable (nonradioactive) isotopes each with a molecular mass of one unit higher, is added to the calibrator series and to the patient samples. For the measurement of methylmalonic acid, for example, a synthetic methyl malonic acid molecule in which three hydrogen atoms have been replaced by deuterium is used as an internal standard; native C12 atoms are often replaced by C13 atoms. The native analyte from the sample and the synthetic internal standard added can be clearly differentiated based on their differing molecular masses using mass spectrometry. The primary readout in the GC-MS analysis is the ratio between the peak area of the SIM track of the native analyte and that of the internal standard. The internal standard labeled with stable isotopes has practically the same physicochemical behavior as the naturally occurring target analyte, which fully compensates for any variations in extraction, derivatization, or ionization between samples.
Isotope dilution GC-MS is a suitable method which qualifies it in almost all areas of laboratory medicine as diagnostic tests for screening newborns for metabolic problems , identifying microbes from human cultures, and measuring the concentrations of therapeutic drugs in blood /, /.
Stable isotope dilution GC-MS offers maximum analytical accuracy and is a suitable reference method technology for the specification of calibration, control, and inter laboratory survey materials in particular.
MS provides unique capabilities in clinical chemistry reference methods, general unknown screening in occupational, environmental, and forensic medicine, measurements that require maximum analytical accuracy (e.g., plasma oxalic acid for the diagnosis of primary oxalosis or phytanic acid for the diagnosis of Refsum disease) /, /.
Strengths and limitations
Very high specificity, high sensitivity; identification of unknown substances.
In HPLC, the mobile phase is a mixture of aqueous buffer solution and organic solvent while the stationary phase consists of densely packed particles with a well defined surface in a steel cylinder. Chains of hydrocarbons on silica particles are often used (e.g., C18 for chains of 18 carbon atoms). The mobile phase is pumped at high pressure (50–800 bar) through the filled separation columns (length approximately 15 cm, internal diameter 4 mm) at a flow rate of around 1 mL/min. Like in gas chromatography, the sample is injected using a software controlled autosampler, which means that analyses can be run over several hours without additional staff input. A 2–100 μL aliquot of the pretreated sample is transferred to the high pressure liquid stream using a special switching valve in the autosampler and carried to the column for separation. The pressure of the liquid stream decreases during HPLC separation and has returned to normal atmospheric pressure by the time the eluate leaves the column and is detected by the detector. In reversed phase chromatography, which is mainly used for medical analytics, less lipophilic substances are eluted before more liphophilic substances, which interact more intensively with the typical nonpolar stationary phases. The higher the proportion of organic solvent (mainly methanol or acetonitrile), the earlier lipophilic substances are eluted.
Three main methods are used in the clinical laboratory to detect eluted substances: UV-VIS detection, fluorescence detection, and electrochemical detection. The first method uses transmission photometers that work in the ultraviolet and visible wavelengths. A prerequisite for this detection method is the presence of UV active double bonds in the target analyte. Although different substances display characteristic absorption maxima at particular wavelengths, the specificity of UV detection is low, especially at low wavelengths.
In the fluorescence detection method, monochromatic light of a particular wavelength is beamed into the measuring cell and any emitted light of a different, longer wavelength is detected. Fluorescence can occur in molecules that contain conjugated double bonds. A fluorescent group can be added to some analyte molecules by means of chemical derivatization.
In electrochemical measurement, detection is based on the specific redox potential of the target analyte. To do this, a voltage is applied in the measuring cell. If a substance is oxidized or reduced, the resulting current can be measured.
Because all commonly used detection principles require the target analyte to have particular molecular characteristics, they are by no means universal . While UV detection has relatively low detection limit (typical detection limits in the range of mg/L), fluorescence and electrochemical detection methods can display very high detection limit for certain analytes (pg/L range).
Because the specificity of the above detection methods for particular substances is significantly limited, the chromatographic separation of all endogenous and exogenous substances of the target analyte in a medical sample must be complete. Separation can be optimized by means of gradient elution (as opposed to constant isocratic elution). In gradient elution, the ratio of organic to aqueous components in the mobile phase varies during the analysis run. Assessment of peak shape is an unreliable method of evaluating the completeness of separation. A broadening of the peak in a sample compared to the peak found when the pure substance is injected (recognizable by a different ratio of peak area to peak height or by peak shoulders) indicates the presence of an unknown interfering substance that co-elutes with the analyte and renders the measurement unusable.
For the analysis of biological samples using HPLC, specific sample preparation is required in all cases. This preparation is usually aimed at removing macromolecular matrix components (proteins) from the sample since the direct injection of serum would rapidly block the HPLC column. The most basic sample preparation involves simply precipitating the proteins with strong acids or organic solvents (e.g., trichloroacetic acid or acetonitrile). Following high speed centrifugation, a clear supernatant is obtained. During more complex liquid phase extraction, an organic solvent such as ethyl acetate, which floats on top of the aqueous phase, is added to the sample. An emulsion is produced by shaking the sample and lipophilic substances are enriched in the solvent. Following centrifugation, the solvent is drawn off, evaporated to dryness and redissolved in the mobile phase for injection. In this way, it is possible to concentrate lipophilic target analytes and increase the detection limit of the methods.
Solid phase extraction is more commonly used. This involves a separate prior chromatographic separation using a single-use cartridge. Serum or urine is added to the cartridge (typically 0.5 to 1 mL), the analytes bind to the stationary phase of the preparation column, and the hydrophilic matrix is washed out with aqueous solutions. The matrix depleted extract is then eluted into the HPLC sample tube with a small quantity of solvent. Target analytes can also be concentrated using this method. Solid phase extraction methods can be automated; in this case, reusable extraction columns are also used (online solid phase extraction).
Proteins such as hemoglobins or transferrin isoforms can also be separated using HPLC. In this case, defined dilutions or sample hemolysis are required. Specialist manufacturers offer HPLC complete kits for a range of parameters that include separation columns, mobile phases, standards, and controls as well as consumables for sample preparation. Some laboratories also use methods developed in house.
HPLC analyses are mainly used in laboratory medicine to measure analytes for which sufficiently specific immunoassays are not available (e.g., in the ambient point-of-care diagnostics). Ambient ionization MS allows direct chemical analysis of unmodified and complex biological samples .
Although the development of effective immunoassays for a pattern of substances that are determined relatively infrequently is technically possible, it is not lucrative for the diagnostics industry. As a result, HPLC methods are used (e.g., for certain anticonvulsants). For some parameters, both effective immunoassays and HPLC methods are available (e.g., HbA1c and homocysteine). The potentially higher analytical specificity of HPLC methods and the generally low cost of the reagents and consumables used must be balanced against the level of staffing required . The choice of method ultimately depends on the specific circumstances of the laboratory in question.
Strengths and limitations
While the development of immunoassays involves complex test antibody production and is therefore almost completely reserved to the diagnostics industry, HPLC methods can often be established quickly and flexibly in experienced laboratories. The specificity of these methods is generally satisfactory but must not be overestimated.
Compared to immunoassays, HPLC methods usually require significantly more effort and specialist personnel. In addition, with a typical analysis time of around 15 minutes, the sample throughput is significantly limited. For many analytes, in particular hormones, the detection limit is insufficient.
HPLC methods cannot be used for all analytes because particular molecular structures are required for detection.
In contrast to GC-MS, conventional HPLC techniques preclude the use of stable isotope labeled internal standards. In this case, structurally related substances must be used that can be clearly separated from the target analytes using chromatography. This means, however, that their extraction properties can vary significantly, which can affect analytic accuracy.
For a long time, the question of how to combine HPLC separation technology with a mass spectrometry detection method (similar to GC-MS) posed a technical challenge. The HPLC eluate could not be transferred directly into a high vacuum like in GC-MS because evaporation of the mobile phase would generate enormous quantities of gas, which would be incompatible with the ultra high vacuum required for mass spectrometry.
Only with the development of electrospray ionization (ESI) it was possible to combine HPLC and mass spectrometry. In ESI, the target analytes are ionized at atmospheric pressure outside the mass spectrometer. To do this, the mobile phase of HPLC is aerosolized by pumping it through a fine capillary tube with the help of a nitrogen stream (nebulizer gas). The spray capillary has a voltage of around 3 kV; as a result, the aerosol spray is electrically charged. A high flow of hot nitrogen gas is directed at the aerosol (approximately 600 L/m at more than 200 °C; desolvation gas) to facilitate rapid evaporation of the solvent. The heat loss due to evaporation protects the analyte from thermal stress. The diameter of the droplets decreases rapidly and the electrostatic repulsion of like charges in the droplets becomes more powerful than the surface tension. At this point the droplets explode, generating many smaller droplets (Coulomb explosion). Eventually, individual molecules in the droplets become charged and individual ions are emitted from the drops due to the repulsion of like charges (). Ionization takes place in an area of approximately 2 cm at the end of the spray capillary. The most frequently used mode is positive ion mode, in which a proton is transferred to the analyte. However, cluster ions may also form with components of the mobile phase (e.g., ammonium, formiate, or sodium adducts with analyte molecules). The entire ion stream generated by ESI is directed toward the narrow, usually orthogonally oriented opening of the mass spectrometer by means of a counter voltage applied to the capillaries. Ion optics are used to direct the ion stream through an area of relatively low vacuum (around 10–3 torr) to the high vacuum area of the mass spectrometer (around 10–6 torr), which is evacuated by turbo-molecular pumps.
In contrast to the electron impact ionization used in GC-MS, molecular disintegration does typically not take place in ESI. It is a soft ionization technique that can also be used for macromolecules such as proteins and DNA. Such molecules can be ionized multiple times using ESI. Because mass spectrometers operate mainly on the basis of the mass-to-charge ratio (m/z) in a range of up to 2,000 molecules of any size can be analyzed using mass spectrometry with the help of deconvolution software (a molecule with a mass of 15,000 and a charge of 10 has a m/z of 1,500).
In addition to electrospray ionization, atmospheric pressure ionization (API) in the form of atmospheric pressure chemical ionization (APCI) or atmospheric pressure photoionization (APPI) is also used, particularly for the detection of nonpolar analytes.
Tandem mass spectrometry involves the coupling of two quadrupole mass filters with an interposed collision cell. This arrangement and the ability to perform two independent quadrupole scans mean that different operating modes are possible in tandem MS. The standard technique for quantitative analysis is multiple reaction monitoring (MRM).
In multiple reaction monitoring, the ion stream generated using ESI is directed toward the first quadrupole, whose radio frequency pattern is set in such a way that only ions with a specific mass-to-charge ratio can pass through the quadrupole. All other ions are deflected. The molecular ions of a target analyte (parent ion) that are filtered out of the total ion stream in this way enter the collision cell, into which an inert gas such as argon is introduced at an extremely low flow rate. The accelerated analyte ions collide with the argon atoms and disintegrate into thermodynamically favorable fragment ions (daughter ions). The stream of fragment ions is now directed toward the second quadrupole, where a single defined fragment ion of the target analyte is filtered out and eventually reaches the ion detector. In MRM mode, therefore, highly specific substance detection is based on the controlled induction of a physicochemical disintegration process (). Tandem MS systems can alternate between more than 100 mass transfers in cycles of significantly less than 1 second, which allows a large number of substances to be quantified simultaneously in an analytic run of only a few minutes.
Advantages compared to previously used chromatographic procedures
Because of the high analytic specificity of tandem mass spectrometry, background signals are largely suppressed, ensuring good detection limit. Furthermore, the importance of sample preparation and chromatographic separation is minimized in comparison to conventional HPLC detection procedures since the simultaneous elution of multiple substances is not a problem; in fact, it is desirable in the case of the obligatory internal standards required for target analytes (). Nevertheless, after the sample matrix has been removed as part of sample preparation, partial chromatographic separation is often useful in order to elute any remaining matrix before the target analytes, which otherwise could lead to ion suppression during ionization. Automated solid phase extraction with column switching and direct coupling to the MS system by means of a short analytical column is a proven solution .
While GC-MS can only be used for a very limited number of medically relevant analytes (thermostable, volatile or volatilizable substances with a molecular mass of less than 500 Da), soft electrospray ionization coupled to tandem mass spectrometry can be used to analyze practically any substance found in the body using highly specific mass fragmentography, regardless of molecular mass. ESI takes place outside the inaccessible MS vacuum zone; only a clean ion stream reaches the vacuum. In GC-MS, on the other hand, the entire GC eluate reaches the source area, which becomes contaminated by non ionized residual matrix. For this reason, the effort required for device maintenance and sample purification is generally much higher for GC-MS than for LC-tandem MS. Because of its low sample preparation requirements, extensive automation options, typical analysis duration of only a few minutes, and low maintenance requirements, LC-tandem MS is suitable for high throughput analysis with low running costs. Tabletop LC-ESI-tandem-MS systems suitable for routine use have been available since the end of the 1990s.
LC-tandem MS is particularly suitable for the extremely accurate quantification of low molecular weight analytes in biological samples, such as the comprehensive multi analyte quantification required in extended neonatal screening for inborn errors of metabolism . Because internal standardization using stable isotopes is possible, reference methods can be developed on the basis of LC-tandem MS.
Strengths and limitations
Very broad spectrum of potential analytes; can be employed for most low molecular weight biologically relevant substances; methods developed independently of the diagnostics industry. Low running costs with good device utilization, highly cost effective technology, generally very high specificity and measurement accuracy.
However, the specificity of LC-tandem MS must not be considered as absolute. For example, conjugate metabolites of an analyte may disintegrate during ionization, before entering the actual mass spectrometer . The sheer number of substances present in the body and the possibility of multiple ionization during ESI entail the risk of multiple substances having identical MRM transitions. For these reasons, some degree of chromatographic separation is also recommended for most applications of LC-tandem MS.
The potential detection limit of modern immunoassays cannot be achieved using the available systems. For most analytes, LC-tandem MS can achieve a detection limit of 1–10 μg/L. This means that the quantification of peptide hormones and proteohormones is not yet possible in a routine setting. The high specificity of LC-tandem MS, based on exact molecular weights, would also pose problems here for analytes with only minor modifications such as truncation. The quantification of such substances is likely to remain the domain of immunoassays.
Collision induced disintegration mass spectra of LC-tandem MS are determined by far more variables than the electron impact mass spectra of GC-MS. It is therefore difficult in the case of LC-tandem MS to develop spectral libraries for the identification of unknown substances in biological samples; general unknown screening in toxicology and environmental medicine will probably remain the domain of GC-MS.
Time-of-flight mass spectrometry is used to precisely determine the mass of molecules or fragment ions based on their flight time in an electric field. When coupled to ESI or matrix-assisted laser desorption ionization (MALDI) and used with specialized software, TOF mass spectrometry is particularly well suited to the characterization of proteins. As such, it can be considered a key technology in proteomics, the comprehensive characterization of all of the proteins of an organism. Proteomics opens up the possibility of identifying new, diagnostically relevant marker molecules. The possibility of characterizing disease-specific protein expression patterns for diagnostic purposes is also the focus of intensive investigation.
Nina Grünen, Christof Meyer-Kleine, Lothar Thomas
Sequencing technologies are grouped in three generations and becoming ever more important in laboratory medicine.
The first generation of sequencing was developed by Frederick Sanger in 1977. The technology is based on chain-termination method (also known as Sanger sequencing). Sanger sequencing targets a specific region of template DNA using an oligonucleotide sequencing primer, which binds to the DNA adjacent to the region of interest .
In the original version T 7 polymerase and chain-terminating dideoxynucleotide analogs (ddNTPs) were used. The ddNTPs are missing the 3' hydroxyl group that is required for 5' to 3' extension of a DNA polynucleotide chain. DdNTPs work as base-specific termination of primed DNA synthesis and cause the chain to break off during the installation. The method was very labor intensive because a lot of specimen and four reaction approaches per DNA-fragment to be examined were required.
The combination of polymerase chain reaction (PCR) and the use of four differently labeled chain-terminating ddNTP analogs greatly simplified the method. By mixing ddNTPs that have been labeled with different color for each unlabeled dNTPs, and template DNA in a polymerase driven reaction, strands of each possible length are produced. The amount of specimen decreased to about 99% and the needed reactions to 75% with simultananous increase in sensitivity and specificity.
In sequencing according to Sanger only one primer is needed. A mixture of fluorescence labeled ddNTPs and dNTPs in an optimized ratio is used for the incorporation of further nucleotides and the elongation of the DNA chain is prevented with the use of ddNTPs. The ddNTPs prevent further inclusion of nucleotides and result in coincidentally inclusion to a stop of DNA synthesis. This is why the method is also known as chain-termination method.
Fluorescence labeled chemically altered ddNTPs are used in addition. This is why the method is also known as chain-termination method. Sanger sequencing was adopted as the primary technology in the “first generation sequencing” of laboratory and commercial sequencing applications.
For Sanger sequencing the reaction tube contains: primer, polymerase, a mixture of ddNTPs and dNTPs. Each of the 4 ddNTP molecules (adenine, guanine, cytosine and thymin) is coupled with a different fluorescent dye but has no 3' hydroxyl group for coupling to the 5' hydroxyl group of a further nucleotide what is required to lengthen the DNA chain. So there is always a base specific signal when installing the last nucleotide.
- Denaturation: a primer is produced (or commercially available) that specifically binds the target sequence of the template. The primer functions as starting point for registration of the nucleotide sequence of the target region. Denaturation occurs at 95 °C. At this temperature the hydrogen bonds of double stranded DNA probe are transformed to single stranded DNA by dissolving the hydrogen bonds of double stranded DNA.
- Primer annealing: the oligonucleotide primer adheres to single stranded DNA at 50 to 65 °C in dependence of the length and sequence of the bases.
- Elongation: polymerase associates nucleotides to single stranded DNA at 68–72 °C. dNTPs are incorporated into the DNA chain until a ddNTP molecule is accidentally incorporated. The ratio of dNTPs to ddNTPs determines the length of the DNA fragment and the number of cycles.
- Repetition: Per cycle the four terminating analogs (ddNTPs as adenine, guanine, cytosine, thymine) are added so that alternate ending of the DNA strands are ddNTPs of adenine, guanine, cytosine, and thymine.
- Assessment: using capillary gel electrophoresis the DNA fragments are separated, resolved to single-nucleotide differences in size. The chain terminated fragments are detected by their fluorescent labels with each color identifying one of the terminating ddNTPs.
NGS is not a new technology, but a multitude of different methods exist. All have one thing in common: massively parallel sequencing. NGS is a second generation technology with the ability of simultaneous sequencing a lot of DNA fragments . The clinical application has been accelerated in laboratory medicine by /, /:
- An increasing amount of well curated clinical, genetic, and genomic data, and about the precision for medical use
- The guidelines of the U.S. Food and Drug Administration (FDA) for the design, development, and validation of NGS
- The Centers for Medicare and Medicaid Services in actively monitoring the rapid innovation of NGS tests.
NGS is an established test method for germline (inherited) and somatic (acquired) genetic mutations. For detection of germline mutations corresponding diagnostic panels are available. The panels may include targeted panel, whole exome, whole genome, or mitochondrial DNA sequencing. Targeted panel testing is possible for a variety of inherited disorders /, /.
DNA library: must be established specifically for every patient.
Target enrichment: is done by PCR (amplicon based sequencing) or hybrid capture technology with subsequent bridge amplification. A lot of polonies arise.
Sequencing: methods of sequencing are pyrosequencing, sequencing by synthesis, and sequencing by ligation. The type of method depends on the sequencer.
Library preparation refers to the process of preparing DNA for use on a sequencer. Many methods are available in breaking DNA into fragments and adding adaptors to the ends. Adaptors may include universal polymerase chain reaction (PCR) primers, barcodes, and hybridization sequences for identification of patient DNA. After binding of adaptors to patient DNA, the DNA is enriched using bridge amplification and be covalently bound to the matrix /, /.
The library is created using PCR or hybridisation and is amplified using PCR subsequently. The methods of emulsion PCR and bridge amplification are used. In emulsion PCR the course of the chemical reaction takes place on the surface of small beads. In emulsion PCR in a water-oil emulsion is located a small bead, a primer, and a DNA molecule, respectively. Each undergoes enrichment or is sequenced directly for targeted testing, exome analysis or whole genome analysis.
- The concentration of DNA fragments is important. The DNA library fragments and small beads are in such a relationship that theoretically a small bead binds a fragment.
- Enrichment may be performed by PCR or by hybridization to complementary sequences. A successful sequencing is prevented by the absence of a DNA-fragment or the presence of two DNA fragments
- Bridge amplification produces DNA cluster by isothermal PCR. DNA fragments previously hybridized are fixed to the glas plate of a flow cell where the clonal amplification takes place. Here too the exact attitude of DNA level is important to get evaluable results and not waste sequencing capacity.
Pyrosequencing: The technology relies on detection of pyrophosphates (dNTPs) instead of using dideoxynucleotides (ddNTPs) to terminate the chain amplification. Different dNTPs are added for generation of sequences of DNA molecules. After one of dATP, dGTP, dCTP, dTTP will complement to the bases of the template strand and release pyrophosphate (PPi) which equals the amount of incorporated nucleotide. The ATP transformed from PPi drives luciferin into oxyluciferin and generates visible light. Principle of Roche 454 and modified according to Iron torrent (pH change is measured).
Sequencing by synthesis: ddNTPs labeled with different fluorescence dyes are available at the same time. After inclusion takes place pyrophosphate and fluorescence dye is split off and the intensity of the fluorescence dye is captured by a camera. Principle of Illumina.
Sequencing by ligation: In Omics technology octamers (8-mer oligonucleotides) are used . A set of four 1,2 probes (each tagged with a different fluorophore) composed by eight bases is added to the flow cell, competing for ligation to the sequencing primer. The first two positions of the probe encompass a known di-base pair specific to the fluorophore bases 3 to 5 are degenerate bases separated from bases 6 to 8 by a phosphorothiolate linkage .
- Massively parallel sequencing of clonally amplified or single DNA molecules that are spatially separated in a flow cell
- The design is a paradigm shift from that of Sanger sequencing, which is based on the electrophoretic separation of chain-termination products produced in individual sequencing reactions
- In NGS, sequencing is performed by repeated cycles of polymerase-mediated nucleotide extensions or, in one format, by iterative cycles of oligonucleotide ligation
- As a massively parallel process, NGS generates hundreds of megabases to gigabases of nucleotide sequence output in a single instrument run, depending on the platform.
For the platforms described below (Roche 454, Ion Torrent SOLiD, Ilumina) are required different strategies to prepare the sequence libraries as well as to detect the signal and ultimately read the DNA sequence. For the Ion Torrent and SOLiD platforms initially local clonal amplification of the initial library molecules into polonies is required to increase the signal-to-noise ratio. The sequencers are not sensitive enough to detect the extension of one base at the target molecule .
The Roche 454 was the first commercially successful next generation system. Emulsion PCR is used to enrich the primer localized on beads. The beads containing multiple copies of the same primer are then sequentially flowed into picotiter plate wells for sequencing /6/. Sequencing is started using a sequence primer which is fixed to single stranded DNA fragments bound to beads. The 4 different dNTPs are washed over the picotiter plate wells. Every time a nucleotide is incorporated during DNA synthesis it releases pyrophosphate, which is converted to ATP. Signals are produced on the principle of pyrosequencing. In the presence of ATP, luciferase converts luciferin to oxyluciferin to generate light, which is then detected and captured by a coupled-charge device camera.
In Ion torrent technology all steps are arranged on a chip containing a flow compartment, a solid state pH sensor, and micro-arrayed wells. The wells are manufactured using process built on standard Complementary metal-oxide-semiconductor (CMOS) technology.
The procedure is as follows: the fragmented DNA of the template (pattern of DNA fragments) is attached to small beads with specific adapter sequences. Amplification of fragmented DNA is carried out via emulsion PCR on the beads with rolling circle replication which produces covalently linked tandem copies of single-stranded DNA, called nanoballs. Next nanoballs are loaded into microwells each binding one nanoball. Generated by the incorporation of each base during DNA synthesis protons are released into a buffer. The change in pH due to proton release, is detected by the sensing layer of the microwell, which translates the chemical signal into a digital one .
Solid means “sequencing by oligonucleotide ligation and detection”. Sequencing by ligation followed by emulsion PCR template (pattern of DNA fragments) preparation is used on the SOLiD platform . Oligo adaptor-linked DNA fragments are coupled with magnetic beads. Each bead-DNA complex is amplified by emulsion PCR. After amplification, the beads are covalently attached to the surface of a glass slide that is placed into a fluidics cassette within sequencer. Two slides are processed per run; one slide receives sequencing reactants as the second slide is being imaged. The ligation based sequencing process starts with the annealing of a universal sequencing primer that is complementary to the SOLiD specific adapters on the library fragments. The addition of a limited set of 8mer semi-degenerative oligonucleotides and DNA ligase is automated by the instrument. When a matching 8mer hybridizes to the DNA fragment sequence adjacent to the universal primer 3' end, DNA ligase seals the phosphate backbone. After the ligation step, a fluorescence readout identifies the fixed base to the 8mer, which responds to either the fifth position or the second position, depending on the cycle number. A subsequent chemical cleavage step removes the sixth through eight base of the ligated 8mer by attacking the linkage between bases 5 and 6, thereby removing the fluorescent group. After wash-out of the unincorporated dNTPs, fluorescence is measured and recorded. Each fluorescence corresponds to a particular dinucleotide combination. Then fluorescence dye is removed and washed and the next sequencing cycle starts .
The Illumina Genom analyzer is based on the concept of sequencing by synthesis with fluorescence detection. The first sequencing step of the Illumina analyzer is to immobilize each DNA fragment and clonally amplify it. The clonal amplification step creates a bead cluster with approximately 1,000 identical copies. Clonal amplification is needed to generate a large enough signal for detection. For immobilization and clonal amplification the Illumina sequencers use sequencing by synthesis with fluorescent detection. All fluorescently labeled nucleotides are added and compete for the next space. The complementary labeled nucleotide will bind but a blocker prevents addition of more than one nucleotide round (reversible terminator chemistry). The remaining nonbound nucleotides are washed away. Laser excitation leads to a fluorescent emission that is recorded simultaneously for each DNA fragment cluster. The fluorescent label and blocker are cleaved, then the next round begins. In each round, one base pair is read from each DNA cluster .
Although the second-generation sequencing technologies deliver a large amount of data with high sensitivity and specificity a new generation is developed: the third generation sequencing . These technologies work on single DNA or RNA molecules without any (pre-) amplification, without use of optical steps, reads of microbites to gigabites in length, no guanine-cytosine coverage bias in DNA sequencing (GC bias) , high read accuracy and would be flexible enough to generate as many sequence reads as are necessary for the specific research question at hand. Therefore the base sequence of DNA polymers can be determined.
The novel technology uses nanopores, tiny biophores with a nanoscale diameter, and measures current changes instead of fluorescence detection approaches . Any particle passing through the pore interrupts the voltage across the channel owing to the nature of the nanopore; each of the four bases leads to a distinctive current change owing to their unique structures. No amplification step or fluorescence labeling is required by this platform and it does not rely on DNA polymerase . Overall, the third generation sequencing technologies provide longer sequence reads, which helps to close gaps in current medical problems.
8. Petersen LM, Martin IW, Moschetti WE,Kershaw CM, Tsongalis GJ. Third-generation sequencing in the clinical laboratory: exploring the advantages and challenges of nanopore sequencing. J Clin Microbiol 2019; 58 (1): e01315–19.
- Targeted types of analysis: this type of metabolomics quantifies analytes but only looks for known or expected analytes related to particular disease. In clinical laboratories indications for targeted types of testing are inborn errors of metabolism e.g., disorders of ammonia detoxification, cholin metabolism, creatin metabolism, and amino acid transport .
- Untargeted metabolomics: a broad spectrum of low molecular weight biomarkers are detected, e.g., for the evaluation of disease or a pattern of low molecular weight analytes. Untargeted metabolomics is typically non quantitative, however a multitude of analytes are measured, which do not contribute to the clinical question.
Metabolomic platforms are based on separation techniques such as ultra-high performance liquid chromatography coupled to tandem mass spectrometry.
Serum, urine, less common cerebrospinal fluid.
Untargeted metabolomics can simultaneously identify a broad range of biochemical analytes in a sample. Metabolomic databases help to guide researchers, but a reference standard is always needed for metabolite identification .
(+) Expression on subpopulation only, under pathological conditions, or depending on the clone. FS, forward scattered light, SS, side scattered light
Figure 52.1-2 An immunoglobulin molecule has a number of idiotopes (epitopes present in the variable region of an immunoglobulin). The idiotopes together make up the isotype of the immunoglobulin. The paratope is a component of an idiotope. Modified from Ref. .
Figure 52.1-3 Principle of monoclonal antibody production. The following steps are necessary: immunization of the animal, splenectomy, and fusion of the splenic lymphocytes with the myeloma cells (hybridization). Separation and cultivation of the hybrid cells in micro cultures. Selection of the Ig-producing hybrid cells. Cultivation of hybrid cell clones that produce the desired antibody (cloning). Production of specific antibodies in larger quantities, either in the ascites of the mouse or in the in- vitro cell culture. Abbreviation: PEG, polyethylene glycol containing medium.
Figure 52.1-5 Precipitin curve: variation in the quantity of precipitate at increasing antigen concentrations in the presence of a constant antibody concentration (diagram according to Heidelberger and Kendall).
Figure 52.1-8 Simultaneous competitive assay. A constant quantity of radio labeled antigen (Ag*) and a variable quantity of analyte antigen (Ag) are allowed to compete for a limited quantity of antibody (Ab). The % B is the percentage binding of Ag* by the antibody; log [Ag] is the logarithm of the analyte antigen concentration in the specimen.
Figure 52.1-9 ELISA: Capture antibodies (Ab) directed against the analyte antigen (Ag) are bound to the surface of a solid phase, e.g. the tube wall. When the sample is added, the analyte antigens are completely bound to the capture antibodies. An enzyme-labeled second antibody (Ab*) is added to label the bound analyte antigen. The labeled antibody binds to the immune complex (capture antibody and analyte antigen) with the formation of a sandwich. For a specific concentration range, there is a linear relationship between antigen concentration and enzyme activity (∆E). This method is also known as a two-site (sandwich) immunoassay.
Figure 52.1-10 Antibody capture assay. This procedure is used if the analyte is an antibody. The corresponding capture antigen is bound to the solid phase. Labeled anti-human globulin is added to measure the analyte, which is bound to the solid phase as an immune complex. The anti-human globulin binds to the analyte with the formation of a sandwich.
Figure 52.1-11 μ-capture assay. Determination of antigen-specific IgM-antibodies. Animal IgG (IgGFc) directed against the Fc fragment of human IgM is bound to a solid phase. The IgM binds to IgGFc. The resulting immune complex is labeled by adding homologous antigen (Ag). The bound antigen is measured by adding homologous labeled IgG antibody (Ab*), which binds to the antigen with the formation of a sandwich.
Figure 52.1-12 Enzyme labeled antigen assay (ELA). Determination of antigen specific IgM. Animal IgG (IgGFc) directed against the Fc fragment of human IgM is bound to a solid phase. The IgM binds to IgGFc. In the next step antigen of the patient sample binds to IgM. The bound antigen is measured by adding homologous labeled IgG antibody (Ab*), which binds to the antigen with the formation of a sandwich (not shown in the figure).
Figure 52.1-16 Autoantibody interference in competitive immunoassay. A) Analyte and labeled antigen (tracer)compete for a limited number of antibodies. B) Thyroid hormone autoantibodies (SHAA) bind both analyte and labeled antigen, thereby producing a falsely low hormone concentration.
A) The capture antibody coupled to the solid phase binds the antigen. The immune complex thus formed is labeled by means of a labeled second antibody.
B) Heterophilic antibodies or human anti-mouse antibodies (HAMA) interfere as follows:
– Anti-idiotype specificity (B, top). A false negative result occurs because the capture antibody is blocked.
– Anti-isotype specificity (B, bottom). A false positive result occurs because more binding sites are created for the labeled second antibody.
C) Crossreactive substances bind to and block the capture antibody (C, top) or the labeled second antibody (C, bottom) to produce a false negative result.
Figure 52.2-2 Illustration of flow cytometric cell analysis. The fluorescence signals generated by cells passing through the laser beam are multiplied by photomultiplier tubes and converted into electrical impulses. These impulses are assigned a channel number depending on the signal intensity. Histograms provide a unidimensional representation of the frequency distribution of a signal across a cell population. Dot plots are used to depict two correlated signals in the form of an x/y diagram.
Figure 52.3-1 Principle of the polymerase chain reaction (PCR). A) Denaturation of double stranded DNA by heating to approximately 95 °C. B) Hybridization of the complementary primers (annealing) at 50–60 °C. The annealing temperature must be determined individually for each primer. C) Elongation of the primers by DNA polymerase at approximately 72 °C. Elongation is terminated by repeated denaturation, followed by annealing and elongation. In the first cycle, fragments of different lengths are produced. As of the second cycle, the counterpart primers hybridize to form amplicons of the required length only. In theory, the number of DNA strands doubles during each cycle (D and E).
Figure 52.3-2 Principle of bisulfite PCR: A) Treatment of single stranded DNA with bisulfite leads to the deamination of cytosine to uracil. In the subsequent PCR, uracil is amplified as thymidine. Methylated cytosine is not affected by bisulfite conversion. B) In pyrosequencing, the sequence of thymidine and cytosine is determined at the position of a CpG dinucleotide. The software evaluates the height of the peak for each nucleotide as a percentage. The thymidine peak corresponds to the degree of methylation within the analyzed CpG dinucleotide. The CpG dinucleotides are regions of DNA where a cytosine nucleotide is followed by a guanine nucleotide in the linear sequence of bases along its 5' to 3' direction.
Figure 52.3-3 Characterization of PCR fragments in agarose gel. The 212 bp long PCR product of the hemochromatosis gene was displayed in 2.0% agarose gel after staining with ethidium bromide. The clinically relevant C282Y mutation can be detected by means of digestion with the restriction enzyme Rsa1. The wild type allele contains a restriction site that results in a 140 bp and a 72 bp fragment after enzyme digestion. In the mutated gene, a second restriction site is present. In this case, the 140 bp fragment is shortened to 110 bp. Lane 1, homozygous mutation; lane 2, heterozygous mutation; lane 3, homozygous wild type; lane 4, uncut PCR fragment; lane 5, marker.