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To achieve accurate and precise results, you need antibodies that consistently bind specifically and selectively to the intended target. Antibody validation must be application-specific to be effective and information on which applications an antibody has been validated in can be found in the Tested Applications section on any antibody datasheet.
Here you will find more information on what is carried out during our application-specific validation processes.
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Developing sandwich ELISAs requires careful selection of a matched antibody pair (a detector and capture antibody) that both bind specifically to the target protein. Screening performance of these antibodies as a pair is crucial for the validation of these antibodies. Only antibody combinations that show high specificity, selectivity, and consistent linearity of dilution for the target are used for further development.
We validate the performance of each antibody pair in plasma, serum, or tissue lysates, using spike-recovery experiments to validate antibody selectivity. This method involves ‘spiking’ purified recombinant target proteins into the biological matrix, which should be recovered and detected within -/+ 20% variation (80–120%) of the kit’s expected protein standard signal in the provided diluent. The recovery observed for the spike should be almost identical in both the biological matrix and the standard diluent for a sample matrix to be considered valid for our ELISA kits.
Linearity studies of sample dilutions are also carried out using sandwich ELISA to ensure that our antibody pairs recognize not only purified recombinant protein, but also native target protein. For this, we measure a standard protein and the native signal of a protein in over a series of dilutions in parallel. If both the standard protein and native signal dilute proportionally, interpolated sample values will have an identical value at all doses tested once corrected for dilution. The concentration of the target protein is determined by multiplying the dilution factor by the calculated concentration (Figure 1).
Figure 1. Linearity of dilution for human PSA SimpleStep® ELISA kit in quantifying human prostate-specific antigen (PSA) in cell culture, serum, plasma, and urine. Diluted samples were measured in duplicate, and the calculated concentrations were multiplied by the dilution factor to determine the final concentration.
To further verify antibody specificity for use in ELISA, we test to what degree the antibodies bind to related proteins or family members. This allows us to gauge any cross-reactivity and interference. For example, to confirm the specificity of our CXCL2 antibodies for use in ELISA, we test using CXCL3 and CXCL1, which share 82% and 63% amino acid identity with CXCL2, respectively. In this case, the related family member proteins show similar to background absorbance values. To determine species reactivity, we will also check the same protein obtained from different species, ie mouse, rabbit, goat, rat, etc to confirm that the antibody is specific for the human protein. Acceptable results for us will show less than 5% cross-reactivity.
Recombinant technology is used to produce the antibodies for our antibody pairs and ELISA kits. This means that once developed, our antibody pairs and ELISAs show high batch-to-batch consistency.
Antibodies are validated in western blot using lysates from cells or tissues that we have identified to express the protein of interest. Once we have determined the right lysates to use, western blots are run and the band size is checked for the expected molecular weight. We will always run several controls in the same western blot experiment, including positive lysate and negative lysate (if possible, Figure 2).
When possible, we also include knock-out (KO) cell lines as a true negative control for our western blots. We are always increasing the number of KO-validated antibodies we provide. In addition, we run old stock alongside our new stock. If we know the old stock works well, this also acts as a suitable positive control.
If the western blot result gives a clear clean band and we are happy with the result from the control lanes, these antibodies will be passed and added to the catalog.
Figure 2. All lanes: anti-p53 antibody [DO-1] – ChIP grade (ab1101). Lane 1: wild-type HAP1 cell lysate (20 µg). Lane 2: p53 knockout HAP1 cell lysate (20 µg). Lane 3: A431 cell lysate (20 µg). Lane 4: Saos-2 cell lysate (20 µg). Antibody binding was detected using the goat anti-mouse IgG H&L (IRDye® 800CW) preabsorbed ab216772. Image shows merged signal (red and green). Green: ab1101 observed at 53 kDa. Red: GAPDH loading control, ab181602, observed at 37 kDa using as secondary goat anti-rabbit IgG H&L (IRDye® 680RD) ab216777.
Immunohistochemistry and immunocytochemistry
IHC and ICC determine whether an antibody recognizes the correct protein based on cellular and subcellular localization. Antibody specificity is confirmed by looking at cells that either do or do not express the target protein within the same tissue. Initially, our scientists will review the available literature to determine the best cell lines and tissues to use for validation. We then check the protein expression by IHC/ICC to see if it has the expected cellular localization (Figure 3). If the localization of the signal is as expected, this antibody will pass and is considered suitable for use in IHC/ICC.
We use a variety of methods, including staining multi-normal human tissue microarrays (TMAs), multi-tumor human TMAs, and rat or mouse TMAs during antibody development. These high-throughput arrays allow us to check many tissues at the same time, providing uniformly as all tissues are exposed to the exact same conditions.
We are currently working towards using KO cell lines for our ICC validation.
Figure 3. IHC image of ab205921 staining PD-L1 in human tonsil formalin fixed paraffin embedded tissue sections. The section was then counterstained with hematoxylin, blued, dehydrated, cleared and mounted with DPX.
Peptide array is a very high-throughput method of antibody validation that allows us to test the specificity of our antibodies against over 500 peptides at one time (Figure 4). We predominantly use peptide array when we test our histone modification antibodies as it allows us to check cross-reactivity between different modifications.
We use a liquid handler to perform six serial dilutions of the peptides, which are then printed onto nitrocellulose slides in triplicate and used to assess the binding specificity of an antibody to all these peptides simultaneously.
Each nitrocellulose slide that we run contains several essential positive and negative controls to assess antibody specificity. We run old stock batches alongside new antibody test batches for side-by-side comparison. We also run peptide array validation of our own antibodies alongside external antibody batches from competitors to compare the performance of antibodies.
Figure 4. ab176916 was tested in peptide array against 501 different modified and unmodified histone peptides; each peptide is printed on the array at six concentrations (each in triplicate). Circle area represents affinity between the antibody and a peptide: all antigen-containing peptides are displayed as red circles, all other peptides as blue circles. The affinity is calculated as the area under the curve when antibody binding values are plotted against the corresponding peptide concentration. Each circle area is normalized to the peptide with the strongest affinity.
Dot blot was frequently used to validate our histone modification antibodies before we began to use peptide array. Many of the histone modification antibodies on our website will still contain the dot blot validation information and states that they are suitable for use in dot blot.
The technique uses a similar principle to peptide array, serial dilutions of several peptides are plotted onto nitrocellulose and we check the specificity of our antibody of interest against these control peptides (Figure 5).
Figure 5. Dot blot analysis of histone H3 (tri methyl K9) peptide (Lane 1), Histone H3K9 unmodified peptide (Lane 2), Histone H3 (crotonyl K4) peptide (Lane 3) and histone H3K4 unmodified peptide (Lane 4) labeled using ab176916, followed by goat anti-rabbit IgG H&L (HRP) (ab97051).
We have now steered away from using dot blot in our more recent antibody batches because our in-house peptide array is much more efficient and allows us to validate our antibodies against hundreds of peptides at once. We will still occasionally use dot blot but only as a confirmation of the peptide array data we obtain.
When we validate antibodies for IP, we carry out a standard IP protocol using magnetic beads as we find that these beads give better results in less time. Once we have isolated our protein of interest we run the supernatant alongside the flow-through from the IP on a western blot. We also run a no-antibody and bead-only control for each lysate we test. which we run on the same western blot. When we do this, we are looking for a clean band at the expected protein size from our IP sample. We are also checking the other lanes of the blot to check for non-specific binding that would appear also in our negative controls (Figure 6).
Figure 6. PD-L1 was immunoprecipitated from 0.35 mg of NCI-H1975 (human non-small cell lung cancer cell line) whole cell lysate with ab228415. Western blot was performed from the immunoprecipitate using ab228415. VeriBlot for IP Detection Reagent (HRP) (ab131366), was used for detection at 1/5,000 dilution. Lane 1: NCI-H1975 whole cell lysate 10 µg (Input). Lane 2: ab228415 IP in NCI-H1975 whole cell lysate (+). Lane 3: Rabbit monoclonal IgG (ab172730) instead of ab228415 in NCI-H1975 whole cell lysate (-).
ChIP testing is carried out on our histone modification antibodies. After antibody specificity is tested by peptide array, ChIP is used to check that the protein target complexed with DNA can be pulled down using our antibody.
For the ChIP experiment, we use chromatin extracted from formaldehyde-cross-linked HeLa cells or mouse NIH/3T3 cells. We carry out ChIP using our standard ChIP protocol and check the performance of the pulldown via qPCR using a panel of primers for positive and negative control loci known for each histone modification. This gives us a profile of our antibody binding in active and inactive genomic control regions (Figure 7).
Figure 7. Chromatin was prepared from HeLa (human epithelial cell line from cervix adenocarcinoma) cells according to the Abcam X-ChIP protocol. ChIP was performed with 25 µg chromatin, 2 µg ab45173 (blue), and 20 µL A/G sepharose beads slurry (10 µL of sepharose A beads + 10 µL of sepharose G beads). Rabbit normal IgG was added to the beads control (yellow). The immunoprecipitated DNA was quantified by real-time PCR (TaqMan approach).
To pass an antibody, we check that the profile we get from the ChIP-qPCR matches what we expect, ie our acetylated histone modification antibody binds to active regions on the genome (Figure 7). We also test the pulldown strength of our new stock and compare it to previous batches to make sure that the pulldown capability is consistent between batches.
We use several standard controls in every ChIP validation experiment, including an IgG and bead-only negative control. In addition to the positive and negative loci we use to control our q-PCR, we also use a no-template negative control for every validation experiment.
We are currently further developing our ChIP methods and ChIP-validation protocols and are working to validate many more of the antibodies currently on our catalog.
When validating antibodies for flow cytometry, we are careful to optimize every step of the staining protocol, including fixation, permeabilization, and washing. To do this, our scientists review the available literature to understand which cell types and conditions are best suited to validate specific antibodies.
We include relevant controls, routinely running unstained, positive, negative, isotype, viability, Fc-blocking, fluorescence minus one (FMO), and single-staining controls. For an FMO control, we stain all our samples with fluorescent conjugates except the one that is being tested. This shows the contribution of the other fluorescent conjugates in the signal of the unlabeled channel. This control is important for determining non-specific binding of an antibody.
Isotype controls are a good negative control that allows us to determine background signal from the signal given by specific antibody binding. These controls use primary antibodies matching the isotype of the primary antibody you are validating but which do not have specificity for the target. We also use many KO cell lines in our flow cytometry validation whenever these are available. If an antibody gives a positive flow cytometry signal and passes all of our control experiements we will make this antibody available for purchase and state that it is suitable for use in flow cytometry.
We are addressing the issue of antibody specificity with an ongoing KO-validation program using human KO cell lines, generated from cellular models via CRISPR/Cas9. KO models provide an excellent standard for antibody validation as they represent a true negative control.
Genetic KO models are so powerful because they allow us to understand the function of a gene by observing the loss-of-function phenotype. We have KO-validated hundreds of antibodies and have also removed any unspecific antibodies from our catalog.
Figure 8. Specificity testing of PD-L1 RabMAb (ab205921) by immunohistochemistry – knockout (KO) testing. Strong IHC detection with anti-PD-L1 is seen in human lung adenocarcinoma tumor cell line L2987. PD-L1 gene was edited in L2987 cells using TALEN constructs targeting exon4 of human PD-L1, transcript variant 1 (NM_014143.3) and complete KO confirmed by deep sequencing in clone L2-14. IHC detection is eliminated in the L2987 L2-14 KO cell line.
Figure 9. Specificity testing of PD-L1 RabMAb (ab205921) by flow cytometry – knockout (KO)testing. Strong detection with anti-PD-L1 TALEN constructs targeting exon4 of human PD-L1, transcript variant 1 (NM_014143.3) and complete KO confirmed by deep sequencing in clone L2-14. Cell surface staining is almost eliminated in the L2987 L2-14 KO cell line.
To ensure the same, accurate results can be obtained across batches of the same antibody, we perform consistency tests to assess batch-to-batch variation.
In the case of recombinant antibodies, consistency between batches is very high, meaning you are unlikely to need to perform additional optimization procedures (eg titration experiments) between batches. This may not be the case with other non-recombinant hybridoma-produced monoclonal and polyclonal antibodies where the degree of variation and drift is inherently higher.
When available and suitable for assay development, recombinant monoclonal antibodies are favored and provide the best batch-to-batch consistency (Figure 8 ).
Figure 10. Batch testing of PD-L1 RabMAb (ab205921) by IHC – analysis of CHO PD-L1 expressed cells. A: rabbit IgG, 5 µg/mL no staining. B: anti PD-L1, 2 µg/mL (batch 1). C: anti PD-L1, 2 µg/mL (batch 3). D: anti PD-L1, 2 µg/mL (batch 4). E: anti PD-L1, 2 µg/mL (batch 5). F: anti PD-L1, 2 µg/mL (batch 6). G: anti PD-L1, 2 µg/mL (batch 7.) All batches (1,3,4,5,6,7) showed consistent results.