Bradford Assay Calculator

The Bradford assay calculator turns raw absorbance readings into a finished protein concentration. You enter a set of protein standards (known concentration in µg/mL paired with the absorbance measured at 595 nm), then enter the absorbance of your unknown sample and an optional dilution factor. The calculator fits a straight line through the standards, reports the calibration equation along with its R², slope and y-intercept, and reads your sample back off that line to give a protein concentration in µg/mL.

The Bradford method is one of the most widely used protein quantification techniques in molecular biology and biochemistry. It relies on Coomassie Brilliant Blue G-250 dye, which shifts its absorbance maximum from about 465 nm to 595 nm when it binds protein under acidic conditions. The more protein in the well or cuvette, the more dye binds and the deeper the blue color, so the absorbance at 595 nm rises with concentration. This Bradford assay calculator automates the standard-curve fitting and back-calculation that you would otherwise do by hand or in a spreadsheet, removing a common source of arithmetic error.

Because every spectrophotometer, dye lot and protein behaves slightly differently, you must run a fresh standard curve alongside your samples each time. That is exactly what this tool models: it does not assume a fixed extinction coefficient. Instead it uses your own measured standards to define the relationship between absorbance and concentration for that specific run, then applies it to your unknown.

The Bradford assay calculator fits a straight line to your standards using ordinary least-squares linear regression. For n standards with concentration x and absorbance y, the slope and intercept are computed from the standard normal-equation formulas. The fitted line is the calibration equation displayed as y = slope · x + intercept.

Once the line is fitted, the calculator inverts it to solve for the unknown. Your sample's absorbance is plugged in as y and the equation is rearranged to find the concentration x. If you diluted the sample before reading it, the measured concentration is multiplied by the dilution factor to recover the concentration of the original, undiluted stock.

• Slope = (n · Σxy − Σx · Σy) / (n · Σx² − (Σx)²)
• Intercept = (Σy − slope · Σx) / n
• Sample concentration = (Aₛₐₘₙₚₑ − intercept) / slope
• Original concentration = sample concentration × dilution factor

The calculator also reports the coefficient of determination, R², which measures how well the line fits the standards. It is computed as R² = 1 − (SS_res / SS_tot), where SS_res is the sum of squared residuals between measured and predicted absorbance and SS_tot is the total sum of squares around the mean absorbance.

The calculator ships with a typical six-point bovine serum albumin (BSA) standard set ranging from 0 to 1000 µg/mL. Walking through these defaults shows exactly how the numbers connect. The standards and their 595 nm absorbances are summarized below.

Running least-squares regression on these six points gives a slope of about 0.000939 absorbance units per µg/mL and a y-intercept near 0.0442, so the fitted equation is roughly y = 0.000939x + 0.0442. The fit is excellent: the calculator reports an R² of 0.9997, meaning the line explains 99.97% of the variation in the standards. With a default sample absorbance of 0.450 and a dilution factor of 1, the calculator solves (0.450 − 0.0442) / 0.000939 ≈ 432.2 µg/mL.

The three calibration outputs each tell you something different about the quality of your Bradford assay. Reading them together is the fastest way to catch a bad run before you trust the sample value.

R² describes how tightly your standards fall on a straight line. The calculator color-codes this value: above 0.99 it is shown in green (excellent linearity), between 0.95 and 0.99 in yellow (acceptable but worth checking), and below 0.95 in red (a poor fit you should not rely on). A low R² usually points to pipetting error, a mislabeled standard, bubbles in the well, or readings pushed outside the dye's linear range.

The slope reflects the assay's sensitivity, the change in absorbance per µg/mL of protein. A steeper slope means small concentration differences produce larger, easier-to-read absorbance changes. The slope depends on the dye lot, the standard protein and the path length, which is precisely why a fresh curve is needed each run. The y-intercept represents the absorbance of your blank or zero-protein point; a small positive intercept (as in the default curve) is normal and reflects residual dye absorbance, while a large intercept can signal a contaminated blank.

Bradford results are only as good as the data feeding the standard curve, so a few practical habits dramatically improve accuracy. First, keep your unknown sample's absorbance inside the range spanned by your standards. The regression is a model of the calibrated region; extrapolating beyond the highest standard is unreliable because the dye-binding response flattens at high protein concentrations.

Second, dilute concentrated samples and enter the dilution factor so the calculator scales the answer back to the original stock. If a reading lands above your top standard, dilute two- or five-fold and re-read rather than trusting an out-of-range value. Third, choose a standard protein that resembles your sample; BSA is the conventional choice, but the Bradford dye binds different proteins with different affinities, so a BSA-based curve can over- or under-estimate proteins with unusual amino-acid composition.

Finally, run standards and samples in replicate, subtract a proper reagent blank, and let color develop for the same fixed time before reading at 595 nm. Detergents such as SDS and Triton X-100 interfere with Coomassie dye and can ruin a Bradford curve, so a compatible assay may be the better choice when detergents are unavoidable. Following these steps keeps R² high and the back-calculated concentration trustworthy.