Lowry Assay Calculator

The Lowry assay calculator turns raw spectrophotometer readings into a protein concentration in micrograms per milliliter (µg/mL). The Lowry protein assay, first published by Oliver Lowry in 1951, remains one of the most cited methods in all of biochemistry because of its high sensitivity and reproducibility. It combines two chemical reactions: the Biuret reaction, in which copper(II) ions chelate the peptide backbone under alkaline conditions, and the subsequent reduction of the Folin-Ciocalteu reagent by tyrosine, tryptophan, and the copper-peptide complex. The result is an intense blue color whose absorbance, read at 750 nm (or 660 nm), is proportional to the amount of protein present.

Because the relationship between protein concentration and absorbance is established empirically, you cannot read concentration directly from a single number. Instead you build a standard curve using known concentrations of a reference protein such as bovine serum albumin (BSA), then interpolate your unknown sample against that curve. This Lowry assay calculator automates the entire workflow: it fits a least-squares regression line through your BSA standards, reports the line equation and the coefficient of determination (R²), and back-calculates the concentration of any sample absorbance you enter. If you diluted your sample before the assay, the dilution factor is applied automatically so the final answer reflects the true concentration of your original stock.

This tool is built for researchers, students, and lab technicians who run protein quantification routinely and need an accurate, repeatable way to convert absorbance into concentration without re-deriving the math in a spreadsheet every time. It pairs naturally with the related Bradford assay and BCA assay calculators on this site for cross-checking results.

The calculator follows the same steps a biochemist would perform by hand, but instantly. First, it collects your standard curve data — a set of BSA concentrations paired with their measured absorbances at 750 nm. The default standards span 0 to 400 µg/mL, which covers a typical Lowry working range. Second, it fits a straight line through those points using ordinary least-squares linear regression, producing a slope, an intercept, and an R² value that tells you how well the line describes the data.

Third, it inverts the line equation to solve for concentration. Given your sample absorbance, the calculator computes the concentration that the standard curve predicts. Finally, it multiplies that interpolated value by your dilution factor to report the concentration of the undiluted original sample. The R² readout is color-coded so you can judge curve quality at a glance: above 0.99 is excellent, 0.95 to 0.99 is good, and below 0.95 signals a poor fit that warrants re-running the standards.

Inputs You Provide

• Standard concentrations (µg/mL): Known BSA concentrations, typically 0, 50, 100, 200, 300, and 400 µg/mL.
• Standard absorbances (750 nm): The optical density measured for each standard.
• Sample absorbance (750 nm): The reading from your unknown protein sample.
• Dilution factor: How many times the sample was diluted before measurement (1 if undiluted).

The calculator builds the standard curve with classic least-squares linear regression. The slope and intercept of the best-fit line are calculated from the sums of your standard data, and the unknown concentration is found by rearranging the line equation to solve for x.

The regression coefficients come from these expressions, where n is the number of standards:

• Slope: m = (n·Σxy − Σx·Σy) / (n·Σx² − (Σx)²)
• Intercept: b = (Σy − m·Σx) / n
• R²: 1 − (SS_res / SS_tot)

Once the line y = mx + b is known, the sample concentration is obtained by solving for x and then scaling by the dilution factor.

After the calculator fits your curve, three numbers matter most: the line equation, the R² value, and the back-calculated concentration. The equation, written as y = mx + b, lets you reproduce the calculation independently or document it in a lab notebook. A higher slope means the assay is more sensitive (more absorbance change per unit of protein), while the intercept reflects the background absorbance of the reagent blank.

The R² value quantifies linearity. For a clean Lowry standard series, R² should exceed 0.99. Values between 0.95 and 0.99 are usable but suggest pipetting variability or a standard drifting toward the nonlinear end of the curve. Anything below 0.95 means the fit is poor and your interpolated concentration is unreliable — re-prepare standards and repeat.

Reading Quality Flags

A critical caveat: your sample absorbance should fall within the range of your standards. If the sample reads higher than your top standard, dilute it and re-measure rather than extrapolating beyond the calibrated region, where the Lowry response becomes nonlinear.

The Lowry method is sensitive but also notoriously susceptible to interference, so technique drives accuracy. The reagents must be added in the correct order and timed carefully because the color develops and then slowly fades. Read all samples and standards in the same time window — ideally within 30 minutes of adding the Folin-Ciocalteu reagent — so that every well is compared at the same point in the color-development curve.

Always run a reagent blank (0 µg/mL) to set your baseline absorbance; the calculator uses this point as part of the regression, so an inaccurate blank skews every result. Prepare fresh BSA standards from a reliable stock and pipette them with the same care you give your unknowns. Because reducing agents (DTT, beta-mercaptoethanol), detergents, EDTA, and ammonium sulfate all interfere with the copper or Folin chemistry, remove or dilute these out before the assay, or choose a compatible method. When in doubt, confirm a surprising result with the BCA assay or Bradford assay, which respond differently to interfering substances and protein composition.