The antibody is truly a workhorse reagent in biomedical research. Antibodies are both highly sensitive and highly specific for particular epitopes, which makes them ideal reagents for research applications (Tables 1 and 2). Additionally, modern biotechnology has facilitated the large-scale production of antibodies. In the 1890s, antibodies were first described by von Behring and Kitasato, who determined that inactive toxins elicited protective immune responses against active toxins in animals and that injections of serum from these protected animals could transfer protection to other animals. For this reason, antibodies were initially described as ‘antitoxins’; however, antibodies were later found to have a much wider repertoire of antigen recognition.

According to the simplest definition, an antibody is the soluble form of the B lymphocyte antigen receptor, and antibodies are produced exclusively by mature B lymphocytes. Five antibody isotypes, which are distinguished by immunoglobulin structure, exist in mice and humans and are composed of 2 heavy chains and 2 light chains. These chains are linked together by disulfide bonds that provide a level of flexibility to the overall molecule. The portion of the molecule without light chains is known as the constant or Fc (crystallizable fragment) region; this region is determined by a fixed set of genes and is identical for all antibodies of a particular isotype in a species. The portion that includes both linked heavy and light chains is known as the variable or Fab (antigen-binding fragment) region; the extreme ends of this region contain the hypervariable sites that recognize and bind to the target antigen epitope. The Fab region is also determined by a fixed set of genes, but further somatic mutations are required to generate unique and highly specific hypervariable sites (Figure 1).

[enlarge]

Figure 1. Schematic representation of the molecular structure of an antibody. The red bars indicate disulfide bonds that link the heavy chains to each other and the light chains.

Both monoclonal (homogenous isotype and antigen specificity) and polyclonal (heterogeneous isotype and antigen specificity) antibodies are available from industry vendors and in individual labs. Polyclonal antibodies are isolated from the sera of animals that have been immunized against a target antigen. While polyclonal antibodies are often specific for an antigen, they contain a mix of different antibodies against different epitopes of the same antigen and may also contain antibodies against unrelated proteins, which can affect assay results. Additionally, a polyclonal antibody supply is dependent on the source animal, and thus no two batches of polyclonal antibody against a particular antigen are identical. In contrast, monoclonal antibodies are obtained from hybridomas or made recombinantly from expression vectors, both of which ensure continuous supply of homogenous antibody.

In immunoprecipitation (IP) assays, also known as “pull-down” assays, antibodies are used to label and precipitate or “pull-down” target antigens from a cell lysate, biological fluid or other aqueous solution. The precipitation processes are facilitated by different kind of beads (e.g., magnetic, agarose, sepharose) that bind to the antibody Fc region, which then allows the antibody-antigen complexes to be separated from the total lysate through centrifugation or other mechanical methods (Figure 2).

IP assays are popular in many cellular and molecular biology research applications. At the most basic level, IP can be used to purify the target antigen for further research use. IP is also commonly used to determine interactions between multiple proteins in homeostatic cells or in cells that have been subjected to a particular treatment. In such assays, an antibody against the first protein would be used for the precipitation step, and subsequent assays such as western blotting (see below) would be used to determine whether a second protein was pulled down with the first.

[enlarge]

Figure 2. Procedural schematic of a standard immunoprecipitation assay. Reprinted with permission from Pierce Protein Biology Products, Thermo Scientific.

Enzyme-linked immunosorbent assays (ELISAs) are used to qualitatively and quantitatively analyze the presence or concentration of a particular soluble antigen such as a specific antibody, or peptide in liquid samples, such as biological fluids and cell culture supernatant. ELISA and similar assays like radioimmunoassays (RIA) can also be used to determine the concentrations of the antibodies, especially in the case of auto-antibodies in blood such as IgG and IgM antibodies against the S protein and RBD of the S protein of SARS-CoV-2 [3], autoantibodies against Sox2 and GABAb in Lambert-Eaton myasthenic syndrome patient plasma [4] or in rheumatoid arthritis [5]. These assays make use of the ability of polystyrene plates or others to bind proteins, including antibodies, as well as the particular specificities of antibodies for target antigens. Generally, these assays incorporate a colorimetric endpoint that can be detected via absorbance wavelength and quantitated from a known standard curve of antigen or antibody dilutions. The detetion antibody is often labelled with an enzyme (horseradish peroxidase or alkaline phosphatase), or one of a myriad of fluorescent tags, or an electrochemiluminescent label (as in the case of the Meso Scale Discovery platform) or through an intermediary label like biotin. ELISA assays can be multiplexed on chips such as the IsoPlexis system [6]. There are 3 commonly used ELISA formats (Figure 3).

[enlarge]

Figure 3. Commonly used ELISA assay formats.

The direct ELISA assay is the simplest format. As indicated in Figure 3, the capture substrate is the specific antigen that is being tested, and the enzyme that catalyzes the color-change reaction is conjugated to the antigen detector antibody [7, 8]. This format is often used to test the efficacy of a new antibody against a known target that can be immobilized on the plate, or to compare the concentration of a purified recombinant protein against a set of known concentration standards. One known disadvantage is that the enzyme reaction must be stopped at a predetermined time-point because as time progresses, any quantity of enzyme will eventually catalyze the reaction and challenge the ability to differentiate between samples with a high and low concentration of analytes. This assay format is seldom commercially available.

[enlarge]

Figure 4. Procedural schematic of a standard direct sandwich ELISA.

The Indirect ELISA assay is a variant of the direct ELISA in that the capture substrate is the specific antigen that is being tested; however, the detection step is mediated by a primary antibody and an enzyme-conjugated secondary antibody which is reactive against the primary antibody. Thus, the primary antibody that recognizes the antigen is not labeled. The main advantage to the use of the secondary antibody here is that it can help amplify a weak signal and increase the signal-detection sensitivity. One common use for the indirect ELISA assay is to detect and quantify for a specific antigen in human serum. The primary antibody and secondary antibody in the case of nanobodies can also be pre-incubated to omit the secondary incubation [9]. The omission also applies to all other immuno methods involving incubation of often polyclonal secondary antibodies, such as Western blotting, IHC, and IC.

[enlarge]

Figure 5. Procedural schematic of a competitive inhibition ELISA.

The Sandwich ELISA assay format is so named because the analyte is “sandwiched” between two different antibodies. Note that the capture substrate in this format is not the analyte, but instead a capture antibody (Figure 3). This format is the most commercially available ELISA assay. The capture antibody in sandwich ELISA is often a monoclonal antibody, a feature that helps increase the specificity of the assay and reduce background noise. The sandwich method is more specific due to the use of the dual antibody system (Figure 4). The capture antibody is sometimes injected into a living animal, as in the case of in vivo cytokine capture assay [10, 11].

A variation of sandwich ELISA assay, called Single-Molecule Assay (Simoa), can increase the sensitivity of ELISA assays from 10^-12 to 10^-19M. In Simoa, tiny beads are coated with a capture antibody; each bead is bound to either one or zero target molecule, and individual beads are detected with another antibody (detection antibody) and a labeling enzyme [12]. It has been, for example, use to detect plasma neurofilament light levels in patients with Alzheimer or Parkinson disease [13-15].

The competitive ELISA assay, for example, the serotonin ELISA kit SEU39-K01 from Eagle Biosciences [16], is a unique variant of the direct/indirect format since here too the capture substrate on the well is the specific antigen. However, the antibody against the analyte is first incubated with the sample and allowed to bind and occupy the antibody in the solution, and only then added to the antigen-coated wells. This way, only unoccupied antibodies bind the antigen on the plate, and the occupied are rinsed. Thus, as the antigen concentration in a sample increases, the signal intensity is expected to decrease. In other words, a larger quantity of analyte in a sample results in fewer free antibodies in the solution, and consequently in a smaller number of labeled antibodies bound to the standard on the plate and a less intense signal (Figure 5).

[enlarge]

Figure 6. ELISPOT. A, procedural schematic. B, representative images of ELISPOT wells. Splenocytes from a vaccinated mouse were stimulated with the target antigen or not for 48 hours; cells were removed, and the wells were subsequently stained with detection antibody and colorimetric reagent.

ELISA assays are widely used tests and are relatively inexpensive and straightforward. It is essential, however, to fully understand the assay procedure and the commonly used controls along with their associated assay tests to correctly interpret the results and to be able to address any challenges.

The three likely problems with the ELISA assay are often associated with the plate-wash instrumentation, the buffers, or the procedure. Using as reference the sandwich ELISA assay which is the most commercially available ELISA format, the procedure is outlined here:

The assay begins with adsorption of the anti-target capture to the microtiter plate. Excess antibody is then washed out of the wells, and a blocking buffer is added to prevent further binding of any reagents to the plate. The wells are generally washed between any reagent addition steps. The sample is then added, and any target antigen binds to the capture antibody. A second anti-target monoclonal antibody detector is then added, which also binds to the target antigen. The detector antibody, often containing an enzyme label, is then stimulated by an enzyme substrate producing a colorimetric change in the well, which could easily be measured by a spectrographic instrument. This change is often reported as an optical density (OD) which is proportional to the amount of captured antigen in the sample.

The three common controls that are outlined below are typically included with each ELISA test, and help ensure proper interpretation of the results [17] :

• The Blank control presents wells that are coated with the capture antibody and also blocked with blocking buffer, but sample or detector antibodies are not added. It helps control for any variation of the plate itself to the measured OD, where the expected values should approach zero. Unexpected high ODs in the blank wells may indicate a plate-washer problem or excess substrate.
• The Zero Concentration control contains all the buffers and reagents from each step of the assay, but the sample contains only the sample buffer without the target antigen. This control helps determine the contribution of all the reagents and buffers in the assay signal, and it presents the true “background.” The expected OD values in this control are only slightly higher than the blank.
• The Non-Specific Binding control isolates the performance of the assay reagents to ensure their proper function in the assay. Wells are first blocked as usual, and then in place of the reagents at each step of the assay, blocking or wash buffer is added. In the final steps, the labeled detector antibody is added along with the substrate development. This control helps assess the contribution of the labeled detector antibody to the overall OD signal of the assay, expecting these wells OD to turn slightly stronger over the blank control wells, but not more than the zero-concentration control. Differences in the signal are attributed to the performance of the labeled detector antibody.

The indirect ELISA assay is associated with a variety of false positive, negative and background noise reactions. This is attributed to the source of the test, human serum, which contains a high concentration of antibodies, and is characterized by high inherent binding affinity to solid surfaces [18]. False positive reactions are attributed to non-specific binding of the sample immunoglobulins to target-antigens by protein-protein interactions. False negative reactions are often a result of the blocking agent that is used in the assay. And a background noise reaction may be caused by hydrophobic binding of immunoglobulin components in the sample specimens to solid surfaces, particularly at low sample serum dilutions. It is therefore important to carefully consider the different types of non-specific reactions that are involved in this assay and use the appropriate controls to prevent misinterpretation of serological antibody assay data [18]. Table 3 summarizes the main challenges and associated resolutions.

ELISPOTs are similar to sandwich ELISA assays in that the detected cytokine or other soluble factor is captured by a monoclonal antibody that is bound to the assay plate and recognized by a second monoclonal antibody that is linked to a molecule that facilitates colorimetric detection. However, in ELISPOT assays, the cytokine-producing cells are cultured directly in capture antibody-coated and blocked assay plates that are lined with a protein-binding membrane, which permits the capture of cytokines from individual cells to be collected in one spot. The cells are discarded after a defined length of time in culture, and the remainder of the assay is performed in a manner very similar to an ELISA assay. However, the final colorimetric step is not analyzed by wavelength detection but by visual detection either under a microscope or with a specialized plate reader. A successful ELISPOT assay of cytokine producing cells will result in a large number of distinct colored spots in each well, and each spot should correspond to a single cell. For example, Gil-Cruz C et al measured the production of IFN-gamma by PBMCs with an IFN-gamma ELISPOT kit from MABTECH [22]. ELISPOT assays are particularly useful for the detection of responses from tiny populations of cells such as antigen-specific T cells from a vaccinated mouse that might not be easily detectable by other types of analyses (Figure 6).

Immunosorbent technology can also be used in combination with microarray technology to yield high-throughput functional proteomic arrays. In these assays, glass or polystyrene slides are coated with either capture antibodies or samples (e.g., cell lysates). The former is similar to a classic sandwich ELISA or ELISPOT since the antigen is bound between plate-bound and free antibodies, and the latter is similar to a direct ELISA since the target antigen is bound directly to the slide. The detection antibodies for both are specific for known antigens and are fluorescently tagged. This technology can screen for alterations in protein concentration and activation status relatively quickly. However, for this array type of assays and other types discussed below, such as single-cell western blotting and ChIP-on-Chip, intra-assay spatial variability must be addressed [23, 24].

[enlarge]

Figure 7. Western blotting. A, procedural schematic. B, representative images. CD8+ T cells were stimulated with crosslinked antibodies against the T cell receptor and the costimulatory receptor CD28 for the indicated times before lysis and western blotting with the indicated antibodies.

Immuno-PCR (I-PCR) is a technique that combines the sensitivity of the nucleic acid amplification by PCR with the specificity of the antibody-based assays resulting in an increase of the detection sensitivity. Typically, it is possible to obtain a 100–10,000-fold increase over the detection limit of the ELISA in several applications. I-PCR was first described by Sano et al in 1992 [25]. Despite the potential of this technique, for several years the use of I-PCR in diagnostic and biomedical applications was limited by some problems, comprising the duration of the assay and, as a consequence, the increase of the error rate. Moreover, one of the major drawbacks was the difficulty in linking antibodies to oligonucleotides. Several solutions have been proposed like quick and efficient conjugation methods (e.g., the Thunder-Link® antibody-oligonucleotide conjugation system by Innova Biosciences). In addition, the introduction of multiplexed and high-throughput techniques largely improved the I-PCR, which is now routinely used as a diagnostic test for some bacterial and viral pathogens, and for the detection of tumor markers and food contamination [26]. The combination of quantitative PCR with I-PCR (qI-PCR) allowed the quantification of low-abundant biomarkers in complex biological samples that are difficult to detect by classical immunoassays [27].

Western blotting is a method in which proteins that have been electrophoretically separated on a gel are transferred to an absorbent membrane via an electric charge. Once blotted, the proteins can be detected with labeled specific antibodies.

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) is performed to detect proteins that have been chemically denatured from their original conformation by SDS. However, some specific antibodies will not recognize the target epitope on a denatured protein, and thus native PAGE can be performed in the absence of SDS.

Blotting can be performed via the wet, semi-dry, or dry method. In the wet approach, the gel is sandwiched with blotting membrane and various filters and submerged into a tank filled with a specific transfer buffer, such as the Tris-glycine one. In the semi-dry method, the gel sandwich is wetted with only a small amount of buffer and is enclosed directly between electrode plates. Finally, in the dry system, no buffer is required, and preassembled ready-to-use stacks containing electrodes, buffer matrices, and membrane are used. Although the dry method is faster, the wet method provides more consistent results and requires less troubleshooting, as the semi-dry method requires a careful fitting of all sandwich components. Furthermore, smaller weight proteins may be transferred through the membrane in semi-dry transfers that are allowed to proceed for too long.

Once transferred and blocked to reduce non-specific protein binding, membranes are incubated with a primary antibody that is specific for the protein of interest. Monoclonal antibodies generally have higher specificity; however, many of these antibodies are generated against a peptide fragment rather than a native protein and thus may not be effective in the detection of proteins separated by native-PAGE. Polyclonal antibodies can also be used but can yield higher background readings. Since primary antibodies are often unlabeled, a labeled secondary antibody that is species-specific for the Fc portion of the primary antibody can be used for the detection step. Most often, the secondary antibody is tagged with an enzyme. Enzyme tagged blots can be visualized by incubating the blot in a chemiluminescent enzyme substrate, followed by exposure to autoradiography film. Positively labeled proteins will appear as dark bands on the film (Figure 7).

Dot blotting is similar to western blotting in that proteins are detected on a membrane; however, for dot blots, the proteins have not been separated electrophoretically. Instead, protein-containing samples are applied or ‘dotted’ directly onto membranes. This is not a fully quantitative method, but it can be useful to detect the presence or absence of particular proteins; for example, dot blotting can be used to detect the location of certain proteins in sucrose gradient separations of cell lysates [28].

[enlarge]

Figure 8. Probed isoelectric focusing (PIF). Procedural schematic. From [1], copyright by PNAS.

Proteins in a very small amount (from as few as 25 cells [1] ) are separated through capillary isoelectric focusing and immobilized in the capillary, and subsequently detected through specific primary antibodies and chemiluminescence [1] (Figure 8).

An article published by Hughes et al in the journal Nature Methods highlights a novel and rapid immunoblotting technique for protein analysis [29]. This single-cell (sc) western blotting method incorporates aspects of both fluidics and microarrays. The authors describe the construction of polyacrylamide gel-coated slides into which microwells are stamped, thus allowing cells from an overlaid suspension to settle into the wells at an average density of approximately one cell/well. Once in the wells, the cells are lysed and briefly subjected to electrophoresis. The separated proteins are then crosslinked to the gel (a step that yields superior stability as evidenced by repeated stripping and reprobing of the gel), stained with primary and fluorochrome-labeled secondary antibodies in a manner similar to conventional western blotting, and visualized via fluorescent microscopy. Although this is the first report of this method and issues such as the diffusion of cell lysates from the wells and consequent losses of protein remain to be resolved, this technique represents a valuable step in protein analysis for the following reasons. First, the single-cell or near single-cell nature of the assay avoids the masking of intercellular variability within a macroscopically homogenous cell population. Second, sc western blotting was found to exhibit high sensitivity and specificity, particularly with respect to off-target antibody binding that could not be distinguished using other single-cell protein analysis methods such as flow cytometry or ICC. Third, this technique provides the advantages of antibody and size-based detection associated with western blotting for samples that would otherwise be of too limited a quantity for conventional analysis. Once optimized, this array-based design will allow the application of functional or morphological experiments that are generally analyzed via conventional western blotting to high-throughput screening scenarios, although intra-assay spatial variation must be addressed [24]. For example, Milo single-cell Western Blot from ProteinSimple was used to estimate the percentage of enteroendocrine cells (neuropods) that expressed synapsin-1 [30].

[enlarge]

Figure 9. Procedural schematic of chromatin immunoprecipitation.

Size Exclusion Chromatography-Microsphere-based Affinity Proteomics (SEC-MAP) applies chromatography followed by precipitation of selected proteins by antibody-coated beads [31, 32]. In contrast to flow cytometry, this method allows the simultaneous detection of a large number of proteins. Since the array measures both size and expression levels of the proteins, it can easily detect proteolytic changes in selected proteins. With regard to other advantages of SEC-MAP, the approximation of the molecular size of a selected protein is an important characteristic of SEC-MAP. However, the appropriate choice and validation of the specific antibodies for the array are crucial for getting valuable and definitive data. In addition, SEC-MAP may be used for characterization of antibody clones for immunoprecipitation [33]. The validation of the results obtained using SEC-MAP is usually performed using flow cytometry, Western blotting or quantitative real-time PCR (RT-qPCR). A recent study by Kanderova et al demonstrated that SEC-MAP might be effectively used for the evaluation of diagnostic markers of acute leukemia in bone marrow samples [34].

Chromatin immunoprecipitation, ChIP, was originally developed in the 1980s to evaluate the interactions of RNA polymerase II with its target genes [35]. In this procedure, cells are fixed in formaldehyde or a similar fixative to crosslink cellular DNA and any associated proteins. The fixed cells are subsequently sonicated to fragment the DNA-protein complexes into fragments of 100-300 base pair (bp) and the fragments are then immunoprecipitated according to standard methods with an antibody specific for the protein of interest such as a histone or transcription factor. In the final step, the DNA-protein complexes are dissociated by a reversal of the crosslinks, and the bound DNA fragments are identified by PCR analysis. At the same time, proteins co-precipitated along with DNA can be identified by mass spectrometry-based approaches [36]. Ideally, the standard ChIP method is used to confirm a suspected association between a protein and putative target gene, due to the requirement for specific PCR primer design (Figure 9). Internal standards can be included in the ChIP experiments (internally calibrated ChIP, ICe-ChIP) to assess the quality of antibodies and other parameters [37].

One limitation of ChIP is the possibility that the crosslinking step might alter the target antigen and thus disrupt antibody binding and IP. In such cases, ChIP can be attempted without the crosslinking step; this procedure is known as IP of native chromatin or N-ChIP [35, 38]. Although the elimination of crosslinking can improve antigen recognition, it is generally only useful if the target protein is known to bind strongly to DNA.

More recently, ChIP has been modified for use in high-throughput analyses. For example, ChIP-on-Chip combines the ChIP technique with microarray technology to permit whole-genome screening of fluorescently tagged sequences. In these assays, precipitated DNA and control (usually unprecipitated input) DNA are labeled with distinct fluorochromes and are hybridized to a DNA microarray chip of particular loci or even whole small genome oligos. The microarrays can be analyzed by standard techniques to provide detailed binding site information for the ChIP sample relative to the control DNA.

ChIP-seq combines ChIP with modern high-throughput sequencing technology to facilitate the identification of previously unidentified target genes. Since next-generation sequencing (NGS) can provide high-resolution analyses of large amounts of genomic material, ChIP-seq is the method of choice for whole large genome ChIP analysis [39].

Cross-linking immunoprecipitation (CLIP) is a methodology first developed by Ule et al in 2003 in their study on interactions between the splicing factor NOVA and a neuron-specific RNA-binding protein (RBP) [40]. It is similar to that for ChIP; however, there are a few notable differences. In CLIP, the suggested crosslinking agent is UV irradiation. Sonication is not required due to the shorter length of RNA transcripts, and cells can be lysed in a standard buffer. However, all lysates must be treated with RNase to protect the transcripts of interest. In 2008 CLIP has been coupled to high-throughput sequencing, generating HITS-CLIP, also known as CLIP-Seq [41]. Much as ChIP techniques permit the analysis of DNA-protein interactions, CLIP allows the analysis of RNA-protein interactions, mapping the RNA binding site on a genome-wide scale. HITS-CLIP, in particular, has been widely used to map protein-RNA interaction sites of several splicing factors, such as PTB [42], FOX2 [43], and Argonaute [44]. However, HITS-CLIP presents some drawbacks related to the efficiency of crosslinking and the accurate determination of RBP binding sites. To overcome these issues, in 2010 Hafner et al developed the photoactivatable-ribonucleoside-enhanced crosslinking and immunoprecipitation (PAR-CLIP) [45] and in 2011 Konig et al proposed the individual-nucleotide resolution CLIP (iCLIP) that allows a resolution at a single-nucleotide level in RBP binding sites [46].

RNA immunoprecipitation (RIP) is a methodology similar to ChIP, in which interactions between proteins and specific RNA sequences are characterized [47]. The principal steps include i) the treatment of living cells with formaldehyde to reversible formation of protein-RNA cross-links; ii) the immunoprecipitation of a protein of interest by using a specific antibody; iii) the reversal of cross-links; iv) the recovered and the analysis of associated RNAs by RT-PCR. RIP can be associated with microarray (RIP-chip), as first described in 2006 [48], or NGS techniques (RIP-seq), thus allowing a rapid and accurate in-depth analysis of RNAs and RBPs, key elements of gene expression regulation through several processes as splicing, editing, stability, and translation of RNA.

Electrophoretic mobility shift assays (EMSAs) are performed to determine the affinities of DNA binding proteins for specific DNA sites [49]. In classical EMSAs, radiolabeled DNA fragments with the site of interest are incubated with the proteins of interest. Relative binding affinity is determined by an upward ‘shift’ of the DNA when subjected to gel electrophoresis since the bound protein increases the molecular weight associated with the DNA and thus causes a slower progression through the gel. To improve specificity, an antibody specific for the protein of interest can also be added to the reactions; if bound, the additional antibody will further slow the migration of the DNA-protein complex through the gel in a process that is known as EM supershift [50].

[enlarge]

Figure 10. Procedural schematic and representative image of immunohistochemistry. Mouse myenteric ganglion was immunolabeled and imaged by confocal microscopy. Green, tyrosine hydroxylase labeling. Blue, autofluorescence. Distributed by Swharden under a Creative Commons license.

Immunohistochemistry (IHC) and immunocytochemistry (ICC) are two methods that are routinely used to evaluate both the presence and the location of proteins in situ within tissues and cells. The methods differ mainly in sample preparation; IHC is performed on mounted and fixed whole tissue sections, while ICC is performed on cells that have been removed from the stroma (e.g., blood cells that have adhered to slides through centrifugation with a cytospin or a monolayer cell culture that has grown on a coverslip) or singular embryos [51] (Figure 10). A variation of immunocytochemistry is the in-cell Western assay (ICW) [52]. During an ICW, cells grown on a multi-well plate are fixed and incubated with primary antibodies in the same way as in immunocytochemistry, and the antibody bindings are usually detected through near-infrared fluorescence with conjugated primary or secondary antibodies (such as DyLight® 680 or 800) to avoid the auto-fluorescence of cells and plates. The signal from each well is often normalized with another near infrared dye with a different emission wavelength such as DRAQ5 [53, 54] or CellTag700 [55] for comparison. Only antibodies that have been validated in immunocytochemistry (note: not Western blotting) can be used in an ICW to ensure that the antibodies and the detection are specific.

There are several common fixation methods for tissues and cells intended for IHC or ICC analysis, and the choice of a fixation method is dependent on the type of analysis. Whole tissue samples that will be analyzed by IHC are often fixed in formaldehyde, a semi-reversible cross-linking agent that is generated from paraformaldehyde (for example, MilliporeSigma P6148 [56] ) and can be further diluted to formalin. Although several reports have noted negative effects on samples due to ‘overfixation’ in formaldehyde, other reports claim no such negative effects and instead warn against not allowing enough fixation time for the formaldehyde to cross-link cellular proteins. Formaldehyde fixation of whole tissues or, in some cases, even whole animals is accomplished by submerging the tissue into a working solution of formaldehyde (e.g., 4% v/v in water). Bead formation after formaldehyde fixation can occur in neuron or retina preparations [57], and sucrose can be added to the fixative solutions to prevent its formation [57]. In addition, fixation may also introduce artifacts. For example, the common cell fixation/permeabilization method with 3% paraformaldehyde (PFA) supplemented with 0.5% Triton X-100 mislocated the enzyme α1,2 ER mannosidase 1 (ERManI), a component of the ER quality control pathway, to Golgi [58]. Archival formalin-fixed, paraffin-embedded (FFPE) tissue sections can be boiled in a retrieval solution of Tris-HCl containing 2% SDS and used in western blot analysis [59].

Alcohols, particularly methanol and ethanol, are often used to fix cells for ICC or for applications in which the DNA should not be damaged. Alcohols are not generally recommended for solid tissues, as they are thought not to preserve tissue morphology to the same extent as formaldehyde [60]. Additionally, alcohol fixation can lead to tissue shrinkage. Acetone is less commonly used as a fixative and is recommended for the fixation of snap-frozen tissues, as it can improve epitope detection, or as a secondary step after methanol, fixation [61]. However, acetone fixation can also result in tissue shrinkage. Finally, for applications in which antigen preservation is essential, tissues can be snap-frozen in isopentane that has been chilled with liquid nitrogen and stored at -80ºC until further processing.

Prior to sectioning and staining, tissues must be embedded in a substrate. Paraffin is often used to embed whole tissue sections. As paraffin is hydrophobic, tissues must first be dehydrated through a series of incubations in graded and increasing concentrations of ethanol. The penultimate step before paraffin perfusion is an incubation in xylenes to completely dehydrate the tissues. After the tissues are perfused for a short time with paraffin, they can be placed (usually within a plastic cassette) in a 65% paraffin bath and then into a mold to harden into small blocks that can be sliced on a microtome. Paraffin embedding is popular due to its relative ease, low cost, long-term durability, and availability. However, paraffin embedding has drawbacks, not least of which is the inability to cut very thin (<5 um) sections. For high-resolution light microscopy and electron microscopy, plastic embedding may be a better choice, as it permits the cutting of very thin sections and might help to retain tissue shape. Additionally, plastic may permit more successful embedding and sectioning of very hard tissues, such as bone. However, plastic is less widely-available and more expensive than paraffin and may interfere with some staining protocols. Tissues that have been previously snap-frozen can be embedded in optimal cutting temperature compound (OCT), which is a solution of water-soluble glycols and resins. OCT leaves very little residue and thus provides low background signal, which is ideal for immunostaining of frozen tissues. One significant disadvantage of OCT is its opacity when frozen, which presents difficulties with regard to the correct position of frozen tissues for sectioning.

Although formalin fixation has many advantages, it can disrupt the 3-dimensional structures of antigen epitopes. Heat-induced epitope retrieval (HIER) can be used on slide-mounted samples to reverse this process. These methods generally employ both heat and an acidic or basic solution; traditionally, the slides are heated in a pH 6 sodium citrate buffer, for example [62], although high-pH buffers are more effective for the retrieval of some antigens. The slides and buffer can be heated in a very hot water bath, a pressure cooker, for example, in an antigen unmasking solution from Vector Laboratories (H-3300) [56], or autoclave, or a microwave, depending on the available equipment. The retrieval process was examined in detail with MALDI-TOF mass spectrometry, and formaldehyde scavengers were found to be novel antigen retrieval agents [63].

Mounted samples can be stained by several methods. Colorimetric staining with enzyme-linked antibodies and colorimetric substrates is commonly used. Enzymes are often alkaline phosphatase (AP) or horseradish peroxidase (HRP). This method is relatively simple, the colorimetric reactions are generally stable, and the slides can be analyzed by standard microscopy. However, endogenous enzymatic activity or non-specific binding of streptavidin-tagged reagents to endogenous biotins can elevate the signal background, and generally, only 1 or 2 antigens can be targeted per sample. Different approaches have been devised to reduce the background and increase the sensitivity. The enzymes and secondary antibodies are sometimes conjugated to an inert polymer such as dextran in a polymer-based detection system [64]. Brown C et al used purple or yellow chromogen in stead of the commonly used 3,3′-diaminobenzidine (DAB) to avoid the anthracotic pigment in cancer cells [65]. In this regard, immunofluorescence, or the use of fluorochrome-tagged antibodies, or through other intermediate steps such as tyramide signal amplification (TSA), for example with TSA detection kits from PerkinElmer [66], is advantageous because it permits the simultaneous labeling and detection of several antigens. However, care must be taken to avoid photobleaching of the linked fluorochromes, which is often irreversible. Integrated systems such as Ventana Discovery XT for immunohistochemistry can also be used to automate and standardize and the process [67, 68]. Integrated image systems such as Xcyto 10 Quantitative Cell Imager from Chemometec can quantify organelle-specific signals for immunocytochemistry, for example, Cas9 nuclear expression [69].

Gold-linked immunostaining (immunogold labeling) is useful for high-resolution analyses. Samples can be incubated with antibodies that are linked to gold particles of varied sizes, permitting the detection of different antigens in a single sample. These particles can be detected with high sensitivity and at high resolution in electron microscope scans, thus permitting a very precise localization of target antigens within cells and tissues. This kind of staining is commonly used to verify subcellular localizations or specific cell-derived structures as exosomes [70].

Multiple antigens on the same sections or samples in IHC and IC experiments can be detected (the multiplex detection) through the use of 1) primary antibodies from different species and their corresponding conjugated secondary antibodies; 2) primary antibodies of different subclasses [71] ; 3) tyramide signal amplification [72] ; 4) cyclic immunofluorescence (t-CyCIF) [73, 74]. The commonly used conjugate HRP can convert a labeled tyramide into a reactive form that covalently binds to tyrosine residues on proteins at or near the HRP and thus enable labeling of multiple antigens through the stripping of primary antibodies. A dedicated article discusses multiplex IHC.

The detection and characterization of the interaction between two proteins are always challenging. Traditional methods such as the two-hybrid assay, are complex and present some limitations. The proximity ligation assay (PLA) is a fast, sensitive and easy technique that allows the simultaneous detection and quantification of protein interactions. Moreover, being an in situ approach, it is possible to determine the cellular localization of the interaction in adherent cell lines and/or frozen or paraffin-embedded tissues. PLA also allows the detection and quantification of single endogenous protein in native conditions. PLA was described for the first time in 2002 by Fredriksson et al [75]. Since then, this technique has become one of the most widely used to analyze interactions in the native protein, under non-perturbed conditions. The PLA methodology is based on the use of two antibodies labeled with oligonucleotides: these antibodies can bind two different epitopes of the same protein or two different proteins. When the antibodies are in close proximity (about 20-40 nm apart), for example when the two analyzed proteins are interacting, the oligonucleotide probes on antibodies will hybridize and bind with two others “connector oligos” to form a continuous circular DNA structure. At this point, a DNA polymerase will amplify this circular structure, which remains covalently attached to one of the PLA antibodies. The amplified DNA molecule can be detected by using standard fluorescent methods, thus directly indicating the interaction between the two analyzed proteins. PLA methodology has been used for a plethora of applications: one of the most commonly used is the validation of biomarkers in the clinical environment [76-78]. PLA has also been used to study the role of specific proteins in biological processes, such as cancer progression. Recently, PLA has been improved to detect post-translational modifications like acetylation, phosphorylation, etc. directly. Hafner AS et al [79] and Brigidi GS et al [80] detected newly synthesized proteins through the use of anti-puromycin antibody, a technique called puro-PLA.

[enlarge]

Figure 11. Scheme of the 96-plex PEA. Adapted from [2].

Following the very same principle of PLA, i.e. dual reporters bearing nucleic-acid extensions that form amplifiable signal when in proximity, the Proximity Extension Assay (PEA) is a sensitive and precise detection technique with high-throughput capabilities in clinical samples. The mechanism of the PEA is the following: a pair of target-specific antibodies carry unique DNA strands that upon binding on the same target hybridise to each other and create a template in a quantitative real-time PCR reaction. Thus, PEA can measure scarce targets, such as cytokines, at a femtomolar range in low volume samples (1 ul) in a quantitative manner [81]. PEA offers great potential in the clinic as it can be easily used in plasma samples [82]. In this study, the authors report IL-8 and GDNF detection in human plasma in a user-friendly, homogenous assay with good recovery, high specificity, and up to 5-log dynamic range in 1 ul sample [82]. Recently, a 96-plex PEA-based immunoassay in body fluids has also been developed [2]. Each of the 96 oligonucleotide antibody-pairs contains unique DNA sequences allowing hybridization only to each other and the amplicons are detected in a high-throughput fluidic chip system. Hence, it has been reported that 92 cancer biomarkers can be detected and quantified simultaneously with high specificity, high sensitivity, and low sample volumes [2].

Interestingly, any polyclonal or any matched pair of monoclonal antibodies can be labeled with unique 40-mer oligonucleotide sequences and become proximity probe sets for the PEA assay [81]. The PEA method offers the advantage that antibody cross-reactivity is hardly detectable and can increase sensitivity of weak antibodies reducing noise signal [2].

[enlarge]

Figure 12. Procedural schematic for surface labeling of cells for magnet-assisted cell sorting and/or flow cytometry. Representative flow cytometric data image shows MACS-sorted CD8+ T cells that were labeled with antibodies against surface CD44 and CD107a.

One of the most common uses for antibodies in both biomedical research and clinical settings is as a means to determine the status of various cell populations by surface labeling and subsequent analysis on a flow cytometer. Surface staining with fluorochrome-tagged antibodies can be performed rapidly on single cell suspensions; this staining tends to be highly specific and can be detected with high sensitivity. Additionally, since many flow cytometers can detect 3 or more fluorochromes simultaneously, cells can be labeled with multiple antibodies, which permits the identification of particular cell subsets as well as surface proteins that are upregulated upon cell differentiation, activation, or apoptosis. Antibodies must be chosen carefully to ensure that the target antigen epitope is expressed on the extracellular surface of the cell, and a blocking agent (e.g., total Ig from the antibody host species) or Innovex #NB309 or BD Biosciences #553142 [83] should be used to avoid nonspecific binding to Fc receptors that are present on several cell types such as various leukocyte subtypes (Figure 11).

Additionally, antibody staining and flow cytometry can be used to detect proteins that reside within the nucleus, cytosol, and endosomes such as transcription factors and cytokines. For this immunostaining application, cells must be fixed and permeabilized with a solution of formaldehyde and a gentle detergent such as saponin, which will reversibly perforate the cell membranes. Antibody staining of intracellular proteins must be performed in the presence of the permeabilizing agent in order to facilitate the transfer of free antibody in and out of the cell. Surface staining should be performed prior to the fixation and permeabilization steps in order to avoid disruptions of surface protein epitopes.

Agonist antibodies specific for cell surface receptors are commonly used to activate immune cells in vitro by binding to and cross-linking the receptors, thus leading to the activation of intracellular signaling pathways. For example, T cells can be stimulated in vitro with a combination of antibodies against CD3, which is closely associated with the T cell receptor (TCR) and must be activated to facilitate TCR signaling, and CD28, which is a costimulatory receptor. However, a second antibody, specific for the Fc portion of the primary antibodies, must be used to crosslink the receptors and induce detectable cell activation fully. Alternatively, the stimulating antibody can be coated onto culture plates prior to the addition of cells (Figures 7, 10) [84].

Antibodies can also be used to block receptors on the cell surface or to neutralize soluble factors in vitro. For example, in helper T cell skewing assays, T cells are cultured in the presence of cytokines that promote differentiation to particular helper T cell subsets, as well as neutralizing antibodies against cytokines that may antagonize differentiation or promote differentiation to a different functional subset. Other modes of antibody functions have been explored as well. For example, Linden JR et al developed rabbit monoclonal antibodies able to block either the binding or the oligomerization of the pore-forming epsilon toxin produced by Clostridium perfringens [85].

In earlier literature such as those from the 1990s, authors used the term 'immunofluorescent staining' to describe flow cytometry.

Fluorescence-assisted cell sorting (FACS) employs the specific binding of fluorochrome-labeled antibodies to cells to sort single cells on the basis of pre-determined fluorescent parameters. This method is very rapid and highly specific; however, specialized flow cytometry equipment is required. Antibodies can also be used to separate or sort cells through binding to magnetic beads in a process known as magnetic-assisted cell sorting (MACS). In MACS, cells are labeled with tagged antibodies that are specific for particular surface markers. The labeled cells are subsequently incubated with very small magnetic beads that bind to the tags. The bead-bound cells can be easily separated from the unlabeled cells by the application of a strong magnet (Figure 11).

Regular antibodies are bulky and can not usually be used to label living cells; however, nanobodies, when conjugated with specific peptides, can enter the cytosol and be used to stain living cells directly [86, 87].

Antibodies can be administered in vivo to deplete specific cell populations for functional analyses. For example, in immunological studies, specific effector lymphocyte subsets can be depleted in mice to determine the consequences of immune responses against specific antigens. Similarly, antibodies can also be used in vivo to neutralize surface receptors on cells or to bind soluble factors, for example, CCL5 [88], similar to the above-described in vitro applications. For these applications, antibodies are generally produced in large quantities from hybridomas to avoid reactions against xenoantigens and are purified to remove cell culture reagents and other potential contaminants [89]. Trim-Away, a protocol for degrading a specific protein, electroporates (or otherwise introduces) an antibody into cells, which, in combination with TRIM21, an E3 ubiquitin ligase and cytosolic antibody receptor, tags the specific protein for proteasomal degradation [90, 91].

Mass spectrometry (MS) is an analytical technique that measures the mass-to-charge (m/z) ratio of product ions to detect, identify and quantify molecules both in simple and complex matrices. Nowadays, MS is an irreplaceable tool for a broad range of fields like proteomics, drug discovery, environmental analysis, biomedical researches. Protein analysis, in particular, has undergone very rapid development since the use of MS instruments, thus allowing an always more accurate and in-depth characterization of such molecules. However, MS has some problems to deal with like the enormous amount of generated data and the presence of high abundance proteins that mask some proteins of interest. The latter point has been partially overcome with the MS targeted approaches, such as multiple/selected reaction monitoring (MRM/SRM). In other words, MS has some trouble with sensitivity while produces highly accurate and specific data. In this framework, the combination of MS with antibody-based affinity methods is a strategy in which a molecule of interest is first captured using immuno-based approaches and then analyzed by MS increasing both specificity and sensitivity. Federspiel JD and Cristea IM provided a detailed immunoprecipitation-based mass spec (IP-MS) protocol for identifying protein interaction [92]. There are several quantitative and qualitative approaches that have been developed by combining immunoassays and MS. The most commonly used are briefly reported below.

MS techniques allow the absolute quantification of a specific protein in a complex sample as plasma. This is achieved by using mixtures of synthetic peptides with heavy isotopes (Absolute QUAntification, AQUA). In 2004 Anderson et al developed the Stable Isotope Standards and Capture by Anti-Peptide Antibodies (SISCAPA) method in which peptides of interest are enriched by using anti-peptide antibodies immobilized on nanocolumns [93]. The protocol is composed by four steps: i) digestion of protein sample with trypsin; ii) addition of heavy isotopes standard peptides like AQUA; iii) immunoenrichment using specific anti-peptide antibodies and iv) peptide absolute quantification by electrospray ionization mass spectrometry (ESI-MS). SISCAPA is used for several applications, for example for the validation and quantification of biomarkers like the human serum transferrin receptor (sTfR) in breast cancer patients [94]. However, the improvement of targeted MS assays directed the studies of immuno-MS towards the combination of SISCAPA and MRM, generating the so-called immuno-MRM.

Multiple reaction monitoring mass spectrometry (MRM-MS) is a targeted quantitative MS approach with high specificity and precision. To increase the sensitivity of this assay, it is possible to enrich the mixture of peptides of interest by immunoaffinity, thus performing an immuno-MRM. This technique is reproducible, can be multiplexed and provides high sensitivity and specificity. The major problem to widely extend the use of immuno-MRM is the lack of validated antibodies specific for this technique. Antibodies are generally produced for the classical immunoassay market (e.g., ELISA, Western blotting), while for the immuno-MRM antibodies should be raised against short, linear, proteotypic peptides. Several studies have been addressed to investigate the use of monoclonal antibodies in immuno-MRM. Unfortunately, monoclonal antibodies are expensive, and their production by hybridoma systems is long. Recently, the feasibility of generating immuno-MRM monoclonal antibodies anti-tryptic peptide antigens by using a recombinant B cell cloning approach has been shown [95].

Immuno-MALDI (iMALDI) is similar to SISCAPA approach: even, in this case, peptides are captured with anti-peptide antibodies, but the enriched sample is then spotted onto a specific target and analyzed by matrix-assisted laser desorption ionization (MALDI) MS. This method has been applied to diagnose different pathologies, such as hypertension [96] and plasma amyloid-beta biomarkers for Alzheimer's disease [97]. However, one of the principal limitations of iMALDI consists in the inability to multiplex the assay.

In the mass spectrometric immunoassay (MSIA), proposed in 1995 [98], a protein sample is incubated with beads coated with a specific antibody and then eluted. The obtained sample can be analyzed directly by MALDI-TOF MS, following a “top-down” approach on the whole protein [99]. The MSIA method can also be coupled to a targeted MS approach like SRM or MRM, by tryptic digestion of eluted sample and subsequent MS analysis. This approach can be automated and multiplexed, as recently reported by Gauthier et al [100].

Mass cytometry combines mass spectrometry and flow cytometry and uses antibodies linked to polymer molecules carrying compounds specific to a selected target [101, 102]. For the analysis, single cells are conjugated with antibodies and undergo the ionization, followed by the evaluation of the ions by time of flight (TOF) mass spectrometry [101]. Recently developed mass cytometry techniques include imaging mass cytometry and multiplexed ion beam imaging. The imaging method uses a laser ablation apparatus to scan and analyze tissue sections after antibody binding [103]. For instance, Hötzel KJ et al used Fluidigm Hyperion imaging mass spectrometer with the ¹⁴⁶Nd-labeled BCL2 EPR17509 antibody to evaluate synthetic antigen gels as practical controls for standardized and quantitative immunohistochemistry [104]. Ion beam imaging applies oxygen primary ions to induce the release of secondary ions from the antibodies bound to the tissue specimens [105]. Both methods can evaluate the relative amounts of the antibodies bound to molecular tissue targets.

Mass cytometry by time-of-flight helps to analyze main cellular processes, such as the cell cycle, differentiation, or hypoxia, activation of signaling pathways and the production of chemokines and growth factors. For instance, this method was effectively applied for the functional analysis of various human T cell subsets, including the spectrum of produced cytokines [106]. Also, the mapping of hematopoietic cells was performed by analyzing multiple parameters using mass cytometry [107]. With regard to experimental design, mass cytometry studies require well-established laboratory protocols. Moreover, the enrichment of target cellular subpopulations prior to the analysis is strongly recommended. Besides, calibration beads are usually suggested to be important for the normalization of the obtained results [108].

Despite the obvious effectiveness of the method, mass cytometry has several limitations, such as difficulty to recover live cells after evaluation due to ionization [109]. Also, it remains difficult to detect molecules with low expression levels.

Biotinylation by antibody recognition is a method to label neighboring moieties, through the non-specific diffusion of the free radicals generated by complexed HRP, which enables the biotinylation of proteins and others [110].

In conclusion, antibodies are an invaluable tool for biomedical research, due to their high sensitivity and specificity, relative ease of production, and flexibility in application uses. The established uses continue to facilitate research, and new developments in antibody-based assay technology are expected to further expand the analytical capacities of basic and translational research laboratories.

Dr. Valeria Severino edited the text and added discussions on PLA, RIP, CLIP, Immuno-PCR, SISCAPA, immuno-MRM, iMALDI, and MSIA on March 10, 2016. Dr. Goldi Kozloski modified the ELISA sections in February 2019. Dr. Nikos Parisis added the section on Proximity Extension Assay in October 2020.