The Bradford assay is a colorimetric method for estimating total protein concentration in a solution. It relies on Coomassie Brilliant Blue G-250 dye, which shifts its absorbance maximum from about 465 nm (the free, reddish-brown cationic form of the dye in acidic solution) to about 595 nm (the blue anionic form) when it binds protein — predominantly at arginine residues, with smaller contributions from lysine, histidine and aromatic residues. Because the shift is proportional to the amount of dye-protein complex formed, absorbance at 595 nm can be converted to protein concentration once the reader has been calibrated against a standard curve of known concentrations, almost always bovine serum albumin (BSA).
This guide walks through preparing a BSA standard dilution series, running the assay, building the standard curve, and working back from a sample’s absorbance reading — through the standard curve and any dilution factor — to a final concentration in mg/mL.
When to use Bradford vs. other protein assays
Bradford is one of several common total-protein quantification methods. Which one fits depends on speed, sensitivity needed, and what else is in the sample buffer. The characteristics below are general tendencies, not fixed values — exact sensitivity and tolerance depend on the specific kit, instrument and sample matrix.
| Method | Typical time | Relative sensitivity | Known interferences | Protein-to-protein uniformity |
|---|---|---|---|---|
| Bradford (Coomassie) | Fast — color develops in 5–10 min, single reagent addition | Moderate; standard and “micro” kit formats cover different ranges | Poor tolerance of ionic/nonionic detergents (SDS, Triton X-100) and strongly basic buffers | Lower — dye response depends heavily on a protein’s arginine/lysine content, so results are reported as BSA-equivalents |
| BCA (bicinchoninic acid) | Slower — typically 30–60 min, often with heat | Comparable to or broader than Bradford depending on kit | Incompatible with reducing agents (DTT, β-mercaptoethanol) and chelators (EDTA) at meaningful concentrations; generally more detergent-tolerant than Bradford | Higher — more consistent response across different proteins |
| Lowry / Folin-Ciocalteu | Slowest — multiple sequential reagent additions and incubations | Historically the reference method; largely superseded | Sensitive to a wide range of buffer components, detergents and reducing agents | Moderate |
| UV absorbance (A280) | Instant — no reagents, non-destructive | Requires reasonably concentrated, pure protein | Nucleic acid contamination inflates readings; needs a known or estimated extinction coefficient for the specific protein | Low unless the extinction coefficient is protein-specific |
Bradford’s main advantages are speed and a single-reagent, single-incubation workflow, which is why it remains common for routine tasks like normalizing loading before a Western blot or an ELISA. Its main limitation is that the dye does not respond uniformly to every protein, so a Bradford result is really a BSA-equivalent estimate, not an absolute measurement of every protein species present.
What you need
- Bradford reagent (commercial dye-based reagent, or dye concentrate diluted per the manufacturer’s instructions)
- BSA standard stock (commonly supplied as a 2 mg/mL solution, or prepared from lyophilized BSA in the same buffer as your samples)
- Diluent that matches your sample buffer as closely as possible, to avoid buffer-mismatch artifacts between standards and samples
- Clean cuvettes or a microplate, and a spectrophotometer or plate reader capable of reading at 595 nm — see UV-Vis spectrophotometer basics and, if it has been a while since the last check, spectrophotometer calibration or microplate reader calibration and maintenance
- Calibrated pipettes across the volume range you’ll be dispensing — small pipetting errors in the standard series are the single biggest source of a poor standard curve
Procedure
1. Prepare the BSA standard dilution series
Dilute the BSA stock into a series of known concentrations that bracket the concentration range you expect your samples to fall in, plus a zero (blank) standard. A typical standard-format Bradford assay is linear across roughly 0.1–1.4 mg/mL protein; “micro” or low-range kit formats extend usable sensitivity down to single-digit µg/mL, at the cost of a narrower upper range. Check the linear range stated for your specific kit rather than assuming the standard-format range applies. A workable general-purpose series, if your kit’s insert doesn’t specify one, is:
| Standard | BSA concentration (mg/mL) |
|---|---|
| S0 (blank) | 0.0 |
| S1 | 0.2 |
| S2 | 0.4 |
| S3 | 0.6 |
| S4 | 0.8 |
| S5 | 1.0 |
| S6 | 1.2 |
Use a fresh serial dilution made up the same day the assay is run — BSA standards do not store reliably once diluted to working concentration. If you need to review how much stock and diluent each step requires, see molarity and solution calculations and buffer and solution preparation.
2. Prepare sample dilutions
If you don’t already have an estimate of your sample’s protein concentration, dilute it at more than one ratio (for example 1:5 and 1:20) so that at least one dilution is likely to land inside the standard curve’s linear range. Record every dilution factor — you will need it in the final calculation step.
3. Add the dye reagent
Add Bradford reagent to each standard and sample well or cuvette. Sample-to-reagent ratios vary by format — cuvette-based assays commonly use on the order of 1 part sample to 20–50 parts reagent, while 96-well microplate formats typically use smaller, more concentrated volumes (for example a few microliters of sample to roughly 200–250 µL of reagent). Follow the ratio specified for your kit rather than a generic figure, since deviating from it shifts the standard curve.
4. Mix and incubate
Mix gently (vortexing can introduce bubbles that distort plate-reader absorbance readings). Incubate at room temperature, protected from direct light, for at least 5 minutes to allow color development to complete. Most standard Bradford protocols report the color as stable for roughly an hour after mixing; beyond that window the dye-protein complex can begin to aggregate or precipitate, especially at higher protein concentrations, which will bias the reading. Read within the stability window your kit’s documentation specifies rather than leaving plates on the bench indefinitely.
5. Read absorbance at 595 nm
Blank the instrument against the S0 (reagent + diluent, no protein) well or cuvette, then read every standard and sample at 595 nm. If any sample reads above your highest standard, don’t extrapolate — dilute it further and re-read, because the Bradford response curve is not perfectly linear once you move past the top of the calibrated range.
Building the standard curve
Plot absorbance (y-axis) against known BSA concentration (x-axis) for each standard and fit a linear regression, y = mx + b, where m is the slope and b is the y-intercept (ideally close to zero if the blank subtraction was done correctly). Check the coefficient of determination (R²); a well-run Bradford standard curve should generally sit at R² ≥ 0.99 across its linear range. If it doesn’t, see the troubleshooting table below before trusting any concentration calculated from it.
Worked calculation (illustrative example)
The values below are a hypothetical, self-consistent data set built to demonstrate the calculation steps — not measured results from any real assay run. Substitute your own instrument’s readings using the same method.
| Standard | BSA (mg/mL) | A595 (hypothetical) |
|---|---|---|
| S0 | 0.0 | 0.005 |
| S1 | 0.2 | 0.120 |
| S2 | 0.4 | 0.235 |
| S3 | 0.6 | 0.350 |
| S4 | 0.8 | 0.465 |
| S5 | 1.0 | 0.580 |
| S6 | 1.2 | 0.695 |
Linear regression on this set gives a slope m = 0.575 A595 per mg/mL and an intercept b = 0.005, i.e. A595 = 0.575 × [protein] + 0.005.
Suppose an unknown sample, diluted 1:10 before the assay was run, gives a reading of A595 = 0.410. Rearranging the regression equation to solve for concentration:
[protein]diluted = (A595 − b) ÷ m = (0.410 − 0.005) ÷ 0.575 = 0.405 ÷ 0.575 = 0.704 mg/mL
That figure is the concentration of the diluted sample that was actually read — and it falls inside the 0–1.2 mg/mL calibrated range, so the reading is valid. Multiply by the dilution factor to recover the concentration of the original, undiluted sample:
[protein]original = 0.704 mg/mL × 10 = 7.04 mg/mL
If a sample’s diluted reading falls above the top standard (here, above A595 ≈ 0.695), the correct response is to dilute it further and re-read — not to extrapolate the regression line past the calibrated range, since the true Bradford response curve tends to flatten (become less sensitive) at higher protein concentrations.
Troubleshooting
| Symptom | Likely cause | Fix |
|---|---|---|
| Standard curve R² below about 0.99, or visibly curved | Degraded or expired reagent; small pipetting errors magnified across the low-volume standard series; standards not made fresh | Prepare BSA standards fresh the same day; use calibrated pipettes appropriate to the volumes involved; if you must extend beyond the linear range, consider a quadratic fit rather than forcing a straight line |
| Sample absorbance reads above the highest standard | Sample is more concentrated than the calibrated range | Dilute further and re-read; don’t extrapolate past the top standard |
| Blank (S0) absorbance is high or inconsistent between runs | Reagent turbidity, contaminated diluent, dirty cuvettes/plate, or expired reagent | Filter the reagent if the kit allows it, use fresh high-purity water, check the reagent’s expiry date, and use dedicated cuvettes/plates for Bradford work |
| Blue staining builds up on cuvettes or plate wells over repeated use | Coomassie dye adsorbs to glass and especially plastic surfaces | Use disposable or dedicated Bradford-compatible cuvettes/plates, and read promptly rather than letting the dye sit |
| Visible precipitate or turbidity in the sample-plus-reagent mixture | Sample too concentrated, or high detergent/salt content driving protein-dye aggregation | Dilute the sample further before adding reagent; if the sample buffer contains incompatible detergents, clean it up (e.g., dialysis or precipitation-based cleanup) before assaying |
| Bradford result disagrees notably with a BCA or Lowry result on the same sample | Protein-to-protein variability — the Coomassie dye’s response depends on a protein’s arginine/lysine content, which differs from BSA | Treat the Bradford number as a BSA-equivalent estimate rather than an absolute value; if precise absolute quantification matters, cross-check with an orthogonal method or a standard protein closer in composition to your sample |
| Little to no color change across the whole standard series | Reagent not added, added in the wrong order or volume, or reagent improperly stored (Coomassie reagent is light-sensitive) | Confirm the reagent-addition step, check storage conditions and expiry, and re-run with a fresh reagent aliquot |
| Readings drift when re-measured later in the same run | Plate/cuvettes read outside the reagent’s stated color-stability window | Read within the incubation window specified for your kit; don’t leave plates sitting on the bench well past that window before reading |
Limitations to keep in mind
- Protein-to-protein variability. Because the dye binds preferentially to basic and aromatic residues, two proteins at the same true mass concentration can give different Bradford readings. Every Bradford number is technically a “BSA-equivalent” estimate, not an absolute mass concentration, unless you calibrate against a standard closer in composition to your actual protein of interest.
- Detergent and buffer sensitivity. Ionic and nonionic detergents (SDS, Triton X-100) and strongly basic or acidic buffers can interfere with dye binding or cause precipitation. If your samples must be lysed or stored in detergent-containing buffer, check compatibility before relying on Bradford, or clean the sample up first.
- Non-linearity at the high end. The dye-binding response tends to flatten above the top of the calibrated range rather than staying linear, which is why diluting an out-of-range sample and re-reading is preferred over extrapolating the regression line.
Frequently asked questions
What is the linear range of the Bradford assay?
It depends on the kit format. Standard-format Bradford assays are commonly linear across roughly 0.1–1.4 mg/mL protein; low-range or “micro” formats extend sensitivity down into the single-digit µg/mL range but cover a narrower span. Always check the range stated for your specific kit rather than assuming a generic figure applies.
Why is my Bradford standard curve not linear?
The most common causes are degraded or expired reagent, small pipetting errors in the low-volume standard dilutions, or standards that weren’t prepared fresh. See the troubleshooting table above.
Can I use the Bradford assay on samples containing SDS or Triton X-100?
Not reliably at meaningful concentrations — the Bradford reagent has poor tolerance for ionic and nonionic detergents. If your lysis buffer contains detergent, consider a detergent-compatible assay format, or clean up the sample (for example by dialysis or precipitation) before running Bradford.
How long after mixing should I read the plate or cuvette?
After at least 5 minutes of room-temperature incubation to let color development finish, and generally within about an hour, since the dye-protein complex can begin to aggregate or the reading can drift beyond that window. Follow the stability window stated for your specific kit.
Why does Bradford give a different number than BCA for the same sample?
Because the two assays respond to different chemistry — Bradford’s Coomassie dye binds preferentially to arginine and lysine residues, while BCA’s copper-reduction chemistry responds more uniformly across amino acid composition. Neither number is more “true” than the other in an absolute sense; both are estimates relative to whatever standard protein (usually BSA) was used to build the curve.
Related guides
- UV-Vis spectrophotometer basics: how it works and measures concentration
- Spectrophotometer calibration: wavelength and photometric accuracy checks
- Microplate reader calibration and maintenance
- Serial dilution technique: how to calculate
- Molarity and solution calculations for the lab
- Buffer and solution preparation lab guide
- Analytical balance calibration and weighing technique
- ELISA protocol basics: step-by-step overview
- Western blot protocol basics
- Cell culture basics: a beginner’s guide
