Enzyme Activity Calculator
Compute enzyme activity in U/mL from the rate of absorbance change in a spectrophotometric assay using the Beer-Lambert law. One unit (U) is conventionally the amount of enzyme converting 1 µmol of substrate per minute.
Last updated: May 2026
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About this calculator
The calculator computes Activity (U/mL) = (ΔA × 1,000,000) / (ε × L × V), where ΔA is the absorbance change per minute (ΔA/min) measured by spectrophotometer, ε is the molar extinction coefficient of the product or substrate in M⁻¹·cm⁻¹, L is the cuvette path length in cm (usually 1), and V is the sample volume in mL added to the reaction. The 1,000,000 factor converts moles per litre into micromoles per millilitre. The Beer-Lambert relationship A = ε·c·L underlies the formula: solving for concentration gives c = A/(ε·L) in mol/L, and the rate of substrate conversion (dc/dt) is found by replacing A with ΔA/min. Typical extinction coefficients: NADH/NADPH at 340 nm = 6,220 M⁻¹·cm⁻¹ (the most common assay), p-nitrophenol at 405 nm = 18,500 M⁻¹·cm⁻¹, DTNB-released TNB at 412 nm = 14,150 M⁻¹·cm⁻¹. Edge cases: the formula assumes initial-rate conditions (ΔA must be in the linear portion of the progress curve) and no substrate depletion; choose a time window where the reaction is < 10% complete. If ΔA exceeds ~0.1/min the kinetic readings may be near the detection ceiling and saturate the spectrophotometer; dilute the enzyme or reduce sample volume. The /V term in this formula is non-standard — most published protocols multiply by total reaction volume divided by sample volume to convert per-cuvette activity to per-mL-of-enzyme-stock; verify against your lab's specific protocol.
How to use
Example 1 — NADH-coupled assay (e.g. lactate dehydrogenase). ΔA = 0.05 absorbance units/min at 340 nm, path length 1 cm, ε = 6,220 M⁻¹·cm⁻¹ (NADH), sample volume 0.1 mL. Step 1: numerator = 0.05 × 1,000,000 = 50,000. Step 2: denominator = 6,220 × 1 × 0.1 = 622. Step 3: 50,000 / 622 ≈ 80.4 U/mL. Verify: a ΔA of 0.05/min at NADH's ε produces a substrate-conversion rate of 0.05/(6220×1) ≈ 8.04 × 10⁻⁶ M/min = 8.04 µmol/L/min; in 0.1 mL that's 0.804 nmol/min total reaction activity ✓. The 80.4 U/mL reflects activity per mL of original enzyme stock added. Example 2 — p-nitrophenyl phosphate assay (alkaline phosphatase). ΔA = 0.30 at 405 nm, path length 1 cm, ε = 18,500 M⁻¹·cm⁻¹ (p-nitrophenol), sample volume 0.02 mL (small volume, concentrated enzyme). Step 1: numerator = 0.30 × 1,000,000 = 300,000. Step 2: denominator = 18,500 × 1 × 0.02 = 370. Step 3: 300,000 / 370 ≈ 810.8 U/mL. Verify: alkaline phosphatase commercial preparations commonly run 100–10,000 U/mL depending on source, so ~810 U/mL is in the realistic range ✓. The high ΔA (0.30) is near the assay's linear ceiling — for routine quantitation, dilute further so ΔA is in the 0.02–0.15 range.
Frequently asked questions
What is an enzyme unit (U) and how does it compare to katal (kat)?
One enzyme unit (U) is conventionally defined as the amount of enzyme that catalyses the conversion of 1 µmol of substrate per minute under standardised conditions of temperature (usually 25 °C or 37 °C), pH (often the enzyme's optimum), and substrate concentration. The SI-derived unit is the katal (kat) = 1 mol/s; conversion: 1 U = 16.67 nkat, so a typical activity of 1,000 U/mL = 16.67 µkat/mL. Most biochemistry literature and commercial enzyme suppliers still use U (the older unit) because most assays were validated in those units, even though IUBMB recommended katal in 1972. Specific activity is the activity per mg of protein (U/mg), used to track purification — as enzyme is purified, specific activity rises while volume activity may fall. Always cite assay conditions because activity depends on temperature, pH, ionic strength, and substrate concentration; the same enzyme can show 10× difference in activity between two assay protocols.
Why is initial-rate kinetics critical, and how do I ensure I'm measuring it?
Enzyme kinetics are only Michaelis-Menten when substrate concentration is constant and product is negligible — true at the very start of the reaction. Over time substrate depletes (slowing the forward reaction) and product accumulates (slowing it further, potentially reversing it), so the absorbance-vs-time curve becomes non-linear and ΔA/min decreases. The standard rule is to use only the linear portion of the progress curve, typically while less than 10% of substrate has been consumed. Practically: plot absorbance vs time over your measurement window, fit a line through the early data, and use that slope. If the line bends downward immediately, your enzyme is over-concentrated or substrate is too low — dilute the enzyme or raise substrate. If absorbance changes too little over the window, your enzyme is under-concentrated — concentrate or use a longer window. Modern spectrophotometers with kinetic software automatically fit the linear portion and report the rate.
How do I choose the right extinction coefficient for my assay?
The extinction coefficient must match the absorbing species being monitored at the specific wavelength used. Most common cofactor assays use NADH or NADPH at 340 nm with ε = 6,220 M⁻¹·cm⁻¹ — both are colourless when oxidised (NAD⁺, NADP⁺) and absorb strongly at 340 nm when reduced, so the rate of appearance or disappearance directly tracks turnover. Other common values: p-nitrophenol at 405 nm = 18,500 M⁻¹·cm⁻¹ (used for ALP, β-galactosidase); TNB (from DTNB) at 412 nm = 14,150 M⁻¹·cm⁻¹ (used for thiol/SH-group quantitation); ATP/ADP cannot be measured directly by absorbance and require coupled assays. Check Sigma's Enzymatic Assay Procedures or the published method for your specific enzyme. Wavelength matters: ε at 340 nm differs from ε at 320 nm for the same molecule, and using the wrong combination will scale your activity by the ratio of coefficients (often 10–100×). Verify with a known concentration of substrate or product before relying on absolute activity numbers.
What are the common mistakes when calculating enzyme activity?
The biggest mistake is using a non-linear region of the absorbance vs time curve — measuring at the wrong time point can over- or under-estimate activity by 50% or more. The second is mixing units: ε is usually published as M⁻¹·cm⁻¹, but some literature uses mM⁻¹·cm⁻¹ (factor of 1000 different); always check. The third is forgetting to subtract a blank — substrate alone often slowly oxidises or hydrolyses, adding an apparent ΔA that has nothing to do with your enzyme; always run a no-enzyme control and subtract. People also confuse sample volume (enzyme aliquot added) with total reaction volume (cuvette volume after all reagents are mixed) — most published formulas use total volume divided by sample volume as a dilution factor, but this calculator's specific formula uses only sample volume. Pipetting errors at small volumes (<10 µL) can be 10–20% and dominate other error sources. Finally, temperature drift during the assay matters: most enzymes' activity doubles for every 10 °C rise, so a poorly thermostated cuvette gives drifting kinetics.
When should I not use this calculator?
Do not use it for end-point assays that measure total absorbance change after the reaction is complete; those need a different formula (activity = total ΔA × volume × dilution / (ε × time × sample volume)), not ΔA/min. It is not appropriate for fluorescence-based assays (using fluorimeters, not spectrophotometers), where the calibration relates emission intensity to product concentration via a standard curve, not Beer-Lambert. Do not use it for radiometric assays, where activity is calculated from disintegrations per minute and specific radioactivity, not absorbance. It is unreliable when the absorbance change is non-linear (substrate inhibition, product inhibition, enzyme inactivation) — in those cases you need a full kinetic analysis or Michaelis-Menten fit. The /V scaling in this specific formula is unusual; check against your assay's protocol because most published formulas separately account for total reaction volume versus sample volume added. Finally, do not use it for crude lysates with multiple enzymes consuming the same substrate without controls — the activity you measure will be the sum of all enzymes, not the one of interest.