ELISA stands for Enzyme-Linked Immunosorbent Assay. It is a plate-based assay technique that uses antibodies and enzyme-mediated colour change to detect and quantify substances such as proteins, antibodies, hormones, and peptides in biological samples.

The basic workflow involves coating a microplate with an antigen or capture antibody, adding the sample, then detecting bound analyte with an enzyme-conjugated antibody. A chromogenic substrate is added, and the resulting colour intensity — measured as optical density (OD) — is proportional to the amount of analyte present.

There are four main ELISA formats:

Direct ELISA: The antigen is coated directly onto the plate and detected with an enzyme-conjugated primary antibody. Simple but less sensitive.

Indirect ELISA: A primary antibody binds the coated antigen, then an enzyme-conjugated secondary antibody provides signal amplification. More flexible and cost-effective.

Sandwich ELISA: A matched pair of antibodies "sandwich" the target antigen — one for capture, one for detection. This is the most common format for commercial kits and offers the best specificity and sensitivity.

Competitive ELISA: The sample antigen competes with a labelled reference antigen for limited antibody binding sites. Ideal for small molecules and haptens that are difficult to sandwich.

For most quantitative applications, a sandwich ELISA is the best choice — it offers the highest specificity because two antibodies must independently recognise the target. This is the format used in the vast majority of Assay Genie ELISA kits.

If your target is a small molecule or hapten (below ~1,000 Da), a competitive ELISA is typically more appropriate because small molecules lack multiple epitopes needed for a sandwich.

Direct and indirect ELISAs are more commonly used for antibody screening, serology, or when developing custom assays.

Quantitative ELISA uses a standard curve of known concentrations to calculate the exact amount of analyte in your sample. This is the standard approach for most Assay Genie kits and is required for research publications.

Qualitative ELISA gives a simple positive/negative result by comparing sample OD to a threshold (cut-off) value. This is common in diagnostic screening (e.g., detecting antibodies to an infectious agent).

Semi-quantitative ELISA estimates relative concentrations (e.g., low/medium/high) without an absolute value. Useful for initial screening before full quantitation.

Sensitivity (also called the minimum detectable dose or lower limit of detection) is the lowest analyte concentration that can be reliably distinguished from a zero-standard blank. It is typically defined as two standard deviations above the mean blank OD.

Detection range is the span from the lowest to the highest standard concentration on the standard curve. You can only reliably quantify samples that fall within this range.

For example, a kit might have a sensitivity of 4.5 pg/mL and a detection range of 15.6–1,000 pg/mL. The sensitivity tells you it can "see" the analyte at very low levels, but you should only report quantitative values within the 15.6–1,000 pg/mL range.

No — serum and plasma have different protein compositions, and the anticoagulants present in plasma (EDTA, heparin, citrate) can interfere with certain assays. You should always stick to one matrix type within an experiment and never mix serum and plasma samples on the same plate.

Most Assay Genie kits specify which sample types have been validated in the product datasheet. If you need to switch matrix types, we recommend running a spike-and-recovery experiment to validate performance first.

We recommend a maximum of 2 freeze-thaw cycles. Each cycle causes protein denaturation, aggregation, and can release proteases from cellular debris — all of which degrade your analyte and reduce measured concentrations.

The best practice is to aliquot samples into single-use volumes immediately after collection and store at –80°C. This avoids repeated freeze-thaw entirely and preserves sample integrity for the life of your study.

It depends on the expected analyte concentration and the kit's detection range. Many Assay Genie kits include recommended dilution factors in the protocol — for example, serum samples for high-abundance cytokines often require 1:5 or 1:10 dilution.

If your samples are likely to have very high analyte levels, you should dilute to avoid the hook effect (where excess antigen paradoxically reduces signal). If you're unsure, run a pilot experiment with a few samples at multiple dilutions (e.g., neat, 1:5, 1:10, 1:50) to find the optimal range.

Always dilute using the sample diluent provided in the kit, not plain PBS — the diluent is formulated to match the matrix of the standards and minimise non-specific binding.

Yes, most Assay Genie ELISA kits are validated for cell culture supernatants and tissue homogenates in addition to serum and plasma. However, these sample types require some extra care:

Cell culture supernatants: Centrifuge to remove cell debris (300×g for 5 min) and test fresh or store at –80°C. Note that serum-containing media can contribute background — if possible, switch to serum-free media 24 hours before collection.

Tissue lysates: Use a gentle lysis buffer without detergents that interfere with the assay (check kit compatibility). Centrifuge at 10,000×g for 10 min at 4°C to remove particulates. Normalise results to total protein concentration (e.g., via BCA assay).

Haemolysed samples are problematic for ELISA. Released haemoglobin absorbs light at 450 nm (the same wavelength used to read TMB-based assays), artificially inflating OD readings. Haemoglobin also has peroxidase-like activity, which can generate false signal with HRP substrates.

If haemolysis is mild (faintly pink), you may be able to proceed with caution — but flag these samples in your analysis. If haemolysis is moderate to severe (red/dark red), re-collect the sample if possible. Ensure gentle blood collection technique and avoid excessive agitation of samples during processing.

Yes — always centrifuge samples before plating, even if they look clear. Particulates, fibrin strands, lipid aggregates, and cell debris can all interfere with antibody binding, cause high background, and increase variability between replicates.

We recommend: 10,000×g for 10 minutes at 2–8°C. Transfer the clear supernatant to a fresh tube before diluting and plating. For plasma samples that tend to form fibrin clots, allow them to sit at room temperature for 15–30 minutes before centrifugation.

It's critical. Antibody–antigen binding kinetics and enzyme reaction rates are temperature-dependent. Cold reagents will produce lower and more variable OD readings, and can cause condensation on the plate that interferes with pipetting accuracy.

Remove all kit components from the refrigerator at least 30 minutes before starting and allow them to equilibrate to 18–25°C. This includes the plate, standards, detection reagents, wash buffer concentrate, and substrate. Gently mix (do not vortex) reconstituted reagents before use.

Washing is arguably the most important step in the entire ELISA protocol. Insufficient washing is the number one cause of high background.

Most protocols specify 3–5 washes per step with 300–400 µL of wash buffer per well. You should:

Aspirate completely between washes — residual liquid dilutes subsequent reagents. Invert the plate and blot firmly on clean paper towels after the final wash. Use fresh wash buffer prepared daily; old buffer can harbour bacterial contamination. Soak for 1–2 minutes per wash if you're experiencing persistent high background.

Not recommended. Each incubation temperature in the protocol has been optimised for the best balance of specific vs. non-specific binding. Increasing the temperature to 37°C when the protocol specifies room temperature will:

Increase non-specific binding (higher background), potentially denature temperature-sensitive antibodies, alter binding equilibrium and reduce specificity, and cause evaporation — particularly problematic for edge wells.

If your time is limited, it's better to plan your experiment around the stated incubation times than to shortcut the temperature conditions.

We strongly recommend using the kit-provided wash buffer. It has been formulated and tested specifically for the antibody pair, blocking conditions, and plate coating used in that kit.

If you must substitute, use 0.05% Tween-20 in PBS (pH 7.2–7.4). However, be aware that the Tween-20 concentration is critical: too little leads to poor washing and high background; too much can strip weakly bound capture antibody from the plate. Commercial formulations may also contain additional stabilisers or preservatives that improve performance.

A dried-out plate is a common cause of high background and irregular results. When the well surface dries, proteins can denature and non-specific binding sites are exposed, leading to patchy and inconsistent signal.

If the plate has dried partially, you can try to rescue it by immediately performing an additional wash cycle (3× with wash buffer, soaking 2 minutes per wash) before adding the next reagent. Results may still be usable, but you should flag these wells and check for unusually high or variable OD values.

To prevent drying: always use plate sealers during incubation, work quickly between steps, and add the next reagent within 1–2 minutes of the final wash aspiration.

Yes — timing consistency is more important than most people realise. If it takes 10 minutes to load an entire 96-well plate, the first wells incubate 10 minutes longer than the last wells. For short incubations (e.g., 30 minutes), this represents a 33% difference in incubation time.

To minimise this effect: use a multi-channel pipette to reduce loading time, add standards and high-priority samples last (so they have the most consistent timing relative to the read), and always add stop solution in the same order you added substrate.

Follow these steps for a reliable standard curve:

1. Reconstitute the lyophilised standard exactly as described in the kit insert. Allow 10 minutes for the protein to fully dissolve, then gently mix by inversion or gentle pipetting — do not vortex, which can denature the protein.

2. Serial dilution: Perform a 2-fold serial dilution series using the provided standard diluent. Use calibrated pipettes and change tips between each dilution step to prevent carryover.

3. Include a zero standard (blank) — this is the diluent alone with no analyte, serving as your baseline reference.

4. Run duplicates at minimum. Triplicates provide better statistical confidence, particularly for the lowest standard points where variability is highest.

5. Prepare standards last, just before plating. Protein adsorbs to tube walls over time, which disproportionately affects low-concentration standards.

Most Assay Genie kits include 7–8 standard concentrations plus a zero (blank). This number is not arbitrary — a 4-parameter logistic (4PL) curve requires sufficient points to accurately define the upper plateau, lower plateau, inflection point, and slope of the sigmoidal curve.

Reducing the number of standards (e.g., to save wells) may seem tempting, but it compromises curve accuracy, especially at the extremes. We recommend always running the full standard range as specified in the kit protocol.

An R² below 0.99 is a warning sign. Before interpreting sample data, investigate:

Check for outlier points: Plot the curve and look for any point that deviates significantly. A single pipetting error on one standard can drag down R². If you identify a clear outlier, you may exclude it with documentation — but never remove points just to improve the fit.

Inspect the low end: The lowest 1–2 standards are the most prone to variability. If only these are off, your mid-range results may still be acceptable.

Re-examine technique: Poor R² often indicates inconsistent serial dilution, reagent degradation, or timing issues. Consider re-running the curve.

Yes — always run a fresh standard curve on every plate. Absolute OD values can vary between plates due to small differences in incubation temperature, wash efficiency, substrate freshness, and plate reader calibration. A standard curve from a previous plate or a different day is not valid for quantifying samples on a new plate.

This also applies if you're running multiple plates from the same kit in a single session — each plate needs its own standard curve.

At minimum, every plate should include:

Blank/zero standard: Diluent only, no analyte. Used to establish baseline OD and subtract background.

Positive control: A sample with a known analyte concentration (often a mid-range standard or a previously validated sample). Confirms the assay is working correctly.

Negative control: A sample known to be free of the analyte. Confirms specificity and that background is low.

For multi-plate experiments, also include a bridging sample — the same sample run on every plate — to monitor inter-assay variability (target CV <15%).

High background is the most common ELISA complaint. The usual suspects:

Insufficient washing: This is the #1 cause. Increase wash cycles to 5×, verify 300–400 µL per well, and consider adding a 1–2 minute soak per wash. Check that your washer isn't clogged or misaligned.

Inadequate blocking: If the plate isn't fully blocked, detection antibodies bind non-specifically to exposed plastic. This is less common with pre-coated kits (like Assay Genie's) but can occur if the plate dried out.

Over-incubation with substrate: TMB continues to develop colour over time. If you exceeded the recommended substrate incubation time, background will be elevated. Add stop solution promptly.

Contaminated buffers: Wash buffer or substrate that has been stored too long or at incorrect temperatures can degrade. Prepare fresh daily.

Cross-reactivity: The detection antibody may be binding something other than the target. Titrate the antibody concentration downward to find the optimal dilution.

If OD is low everywhere — including the highest standards — the issue is assay-wide, not sample-specific. Check:

Did you skip a step? A missed addition of detection antibody, enzyme conjugate, or substrate is more common than you'd think. Work through the protocol with a checklist.

Reagent storage: Were reagents stored correctly? HRP conjugates are sensitive to light and temperature. Check expiry dates on every component.

Sodium azide: If any of your buffers contain NaN₃ (a common preservative), this will irreversibly inhibit HRP. Use azide-free buffers.

Plate reader settings: Confirm you're reading at the correct wavelength — 450 nm for TMB, 405 nm for pNPP. Also check that the reference wavelength (if used) is set to 540–570 nm, not another value.

Wrong antibody pair: Verify that the capture and detection antibodies are specific to your target analyte and the correct species.

High intra-assay coefficient of variation (CV >10%) between replicates is almost always a pipetting issue. Common causes and fixes:

Pipette calibration: Have your pipettes been calibrated recently? Even a 5% volume error significantly impacts results at low concentrations.

Technique: Pre-wet tips before aspirating, pipette slowly for viscous samples (plasma, concentrated protein solutions), and use reverse pipetting for small volumes. Avoid touching the well walls.

Bubbles: Air bubbles in wells scatter light and give artificially high or erratic OD readings. Centrifuge the plate briefly (300×g, 1 minute) after adding samples, or tap the plate gently on the bench.

Multi-channel pipettes: Use multi-channel pipettes for adding reagents and wash buffer to reduce well-to-well timing and volume differences.

Edge effects appear as consistently higher or lower OD values in the outermost wells (Row A, Row H, Column 1, Column 12) compared to interior wells. This is caused by uneven temperature distribution and evaporation — edge wells lose moisture faster and experience slightly different incubation temperatures.

Prevention strategies:

Use adhesive plate sealers for all incubation steps. This dramatically reduces evaporation. Place plates in a humidified incubation chamber rather than on an open bench. Avoid placing plates near draughts, air conditioning vents, or warm equipment. Consider not using outermost wells for critical samples — reserve them for blanks or controls instead. Do not stack plates during incubation.

When standards work fine but samples underperform, the issue is typically matrix-related:

Matrix interference: Sample components (proteins, lipids, heterophilic antibodies) can inhibit antibody–antigen binding. Try increasing the sample dilution — if values rise proportionally, the matrix was suppressing signal at the lower dilution.

Analyte degradation: The target protein may have degraded due to improper storage, too many freeze-thaw cycles, or protease activity. Compare fresh vs. stored samples if possible.

Wrong sample type: Confirm the target is actually expressed/secreted in your biological system. For example, not all cytokines are detectable in all tissue types or time points.

Hook effect: If the analyte concentration is extremely high, it can paradoxically produce low signal in a sandwich ELISA. Test at multiple dilutions (1:10, 1:100, 1:1000) to rule this out.

Two validation experiments can identify matrix effects:

Spike-and-recovery: Add a known concentration of analyte (standard) to your sample, then measure the total. Calculate: Recovery (%) = (Measured / Expected) × 100. Acceptable recovery is 80–120%. If recovery is low, the matrix is suppressing signal; if high, it's enhancing it.

Linearity of dilution: Serially dilute your sample (e.g., 1:2, 1:4, 1:8, 1:16) and measure each. When you multiply back by the dilution factor, all values should agree within ±20%. If they don't — especially if higher dilutions give higher back-calculated concentrations — matrix interference is present at lower dilutions.

The hook effect (also called the prozone effect) occurs in sandwich ELISAs when the analyte concentration is extremely high. At very high concentrations, excess antigen saturates both the capture and detection antibodies separately, preventing them from forming the "sandwich" complex. The result is a paradoxical decrease in signal that can mimic a low-concentration sample.

To detect it: always test suspect samples at multiple dilutions (e.g., neat, 1:10, 1:100, 1:1000). If the measured concentration increases with dilution (after correcting for the dilution factor), the hook effect is present at the lower dilution.

The hook effect is most commonly seen with high-abundance analytes in undiluted samples — this is one reason why kit protocols often specify a recommended minimum dilution factor.

Use a 4-parameter logistic (4PL) regression — this is the gold standard for ELISA data. The 4PL model accurately describes the sigmoidal dose–response relationship by fitting four parameters: minimum asymptote, maximum asymptote, inflection point (EC50), and Hill slope.

Avoid linear regression, which distorts concentrations at both the upper and lower ends of the curve where the dose–response relationship is non-linear. Even log-linear transformations are inferior to 4PL for most ELISA applications.

For asymmetric curves, a 5-parameter logistic (5PL) model may provide a better fit by adding an asymmetry factor. Most ELISA analysis software (including free tools like MyAssays.com) supports both 4PL and 5PL.

Yes — always. Subtract the mean OD of your blank wells (zero standard) from all other wells before plotting the standard curve and calculating sample concentrations. This corrects for non-specific background absorbance from the plate, buffers, and substrate.

If your plate reader supports dual-wavelength reading, also use a reference wavelength (540–570 nm for TMB at 450 nm) to correct for optical imperfections in the plate itself.

Above the highest standard: Re-run the sample at a higher dilution (e.g., 1:10 or 1:100) so the value falls within the standard curve range. Do not extrapolate — the curve becomes unreliable beyond the highest calibrator.

Below the lowest standard: Report as "<LLOD" (below the lower limit of detection) or "<LLOQ" (below the lower limit of quantification). For statistical analysis, common approaches include substituting LLOD/2 or LLOD/√2, but always state your substitution method.

Never extrapolate in either direction. The mathematical model is only validated within the range defined by your standards. Results outside this range have unknown accuracy.

Standard acceptance criteria for ELISA:

Intra-assay CV (between duplicates on the same plate): Should be <10%. Duplicates with CV >10% should be flagged and investigated. If you identify a clear pipetting error (e.g., bubble, missed well), you may use the remaining replicate — but document the reason.

Inter-assay CV (between plates or between runs): Should be <15%. Higher inter-assay CV suggests inconsistencies in reagent preparation, incubation conditions, or plate handling between runs.

Standard curve: R² ≥ 0.99 for a 4PL fit. Individual standard CVs should also be <10%.

Not necessarily. Absolute OD values are instrument-specific and can vary significantly between plate readers due to differences in light source, detector sensitivity, and calibration. The datasheet values are generated under our specific laboratory conditions and serve as a general guide, not an absolute target.

What matters is the shape of your standard curve (sigmoidal), the R² value (≥0.99), and whether your controls fall within expected ranges. If all three criteria are met, your data is likely valid even if the absolute OD numbers differ from the datasheet.

If you're concerned, contact our technical support with your raw data — we'll review it and advise whether a re-run is needed.

Before opening: Store the entire kit at 2–8°C (standard refrigerator) unless the kit insert specifies otherwise. Do not freeze the kit — freezing can damage the pre-coated plate, crack liquid reagent vials, and denature enzyme conjugates.

After opening: Return unused components to 2–8°C immediately. Reseal the plate pouch with the desiccant inside to prevent moisture from degrading the coated surface. Reconstituted standards and reagents should be used within the timeframe specified in the kit insert (typically within 1–7 days when stored at 2–8°C).

We don't recommend it. The expiry date reflects the period during which we guarantee kit performance based on stability testing. Beyond this date, enzyme conjugates lose activity, coated antibodies may degrade, and substrate reagents can oxidise — all of which reduce sensitivity and increase variability.

If you absolutely must test an expired kit (e.g., it's the only option and replacing it would cause a significant delay), run a full standard curve first and include positive controls to assess whether performance is still acceptable. If the curve shape, R², and control values look good, the results may be usable — but document the expired status in your records.

Yes — most Assay Genie kits use breakable strip wells specifically for this purpose. This allows you to use only the number of wells you need and save the rest for later experiments.

After removing the strips you need, immediately reseal the remaining strips in the original foil pouch with the desiccant pack. Store at 2–8°C and use within 1 month of first opening. Always note the date opened on the pouch.

Be aware that you'll still need to run a fresh standard curve each time you use stored strips, as plate performance may shift slightly over time.

Reconstituted standards are generally stable for up to 7 days at 2–8°C or up to 1 month at –20°C when aliquoted. However, this varies by analyte — some proteins are inherently less stable than others.

For best results, prepare standards fresh for each experiment. If you must store reconstituted standard, aliquot into single-use volumes to avoid freeze-thaw degradation, and always run a fresh standard curve alongside to account for any loss of activity.