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The bicinchoninic acid (BCA) assay is a colorimetric method for estimating total protein concentration in a solution. It works in two linked steps: the peptide bonds in a protein reduce Cu²⁺ ions to Cu¹⁺ in an alkaline solution (a biuret-type reaction), and two molecules of bicinchoninic acid then chelate each Cu¹⁺ ion to form an intensely purple, water-soluble complex with an absorbance maximum at 562 nm. Because color intensity scales with the amount of complex formed, absorbance at 562 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). The method traces to Smith et al., Analytical Biochemistry 150(1):76–85 (1985), “Measurement of protein using bicinchoninic acid,” and its two-reagent, single-color-endpoint format is now one of the two most widely used total-protein assays alongside the Bradford (Coomassie dye-binding) method.
This guide walks through preparing the working reagent and BSA standard series, running the assay (including the standard vs. accelerated/micro incubation options), building the standard curve, and working back from a sample’s absorbance reading — through the curve fit and any dilution factor — to a final concentration in mg/mL, plus the interferences that most often produce a bad curve or an implausible sample value.
BCA vs. Bradford vs. Lowry: when BCA is the better choice
BCA and Bradford are the two methods a bench scientist reaches for most often; which one fits depends on what else is in the sample buffer and how much protein-to-protein consistency matters.
| Method | Typical time | Detergent tolerance | Known interferences | Protein-to-protein uniformity |
|---|---|---|---|---|
| BCA (bicinchoninic acid) | 30–60 min at 37°C (standard), or ~30 min at 60°C for an accelerated read | Good — tolerates SDS and most nonionic/ionic detergents at working concentrations | Reducing agents (DTT, β-mercaptoethanol, TCEP) and metal chelators (EDTA, EGTA) at meaningful concentrations; some tolerance of common lipids and membrane components can still shift readings | Higher — more consistent response across different proteins than Bradford |
| Bradford (Coomassie) | Fast — 5–10 min, single reagent addition, read at 595 nm | Poor — ionic/nonionic detergents and strongly basic buffers distort the dye response | Detergents, and buffers far from neutral pH | Lower — dye response depends heavily on a protein’s arginine/lysine content |
| Lowry / Folin-Ciocalteu | Slowest — multiple sequential reagent additions and incubations | Poor | Sensitive to a wide range of buffer components, detergents and reducing agents | Moderate |
The practical rule of thumb: if a lysis buffer contains SDS or another detergent, BCA is usually the safer default. If a buffer contains DTT, β-mercaptoethanol, or EDTA at working concentrations, Bradford (or a reducing-agent-compatible BCA kit variant) is often the better fit — see the interferences section below before assuming either assay will tolerate a buffer as-is. For a full walkthrough of the Coomassie-dye method and its own standard-curve calculation, see the Bradford protein assay guide.
What you need
- BCA working reagent, freshly prepared by mixing Reagent A (bicinchoninic acid, sodium carbonate, sodium bicarbonate and sodium tartrate in alkaline solution) with Reagent B (4% cupric sulfate) — commonly at a 50:1 ratio (A:B), though the exact ratio depends on the kit; mix only as much as needed for the run, since the working reagent has a limited working-day shelf life once combined
- BSA standard stock (commonly supplied as a 2 mg/mL solution, or prepared from lyophilized BSA in the same buffer as the samples)
- Diluent that matches the sample buffer as closely as possible, so standards and samples experience the same background chemistry
- A microplate (96-well is standard) or cuvettes, and a spectrophotometer or plate reader capable of reading at 562 nm — see UV-Vis spectrophotometer basics and, if calibration is due, spectrophotometer calibration or the microplate reader calibration and maintenance guide
- A heat block or incubator set to 37°C (standard protocol) or 60°C (accelerated/micro protocol), unless running the room-temperature 2-hour variant some kits support
- Adjustable pipettes across the volume range the protocol calls for — see pipetting technique best practices if standard-curve replicates are coming out noisy
Step-by-step protocol
1. Prepare the BSA standard dilution series
Make a serial dilution series from the BSA stock down to the diluent blank — a typical standard BCA range runs from roughly 20 to 2,000 µg/mL in 6–8 points, while a micro-BCA kit variant covers a much lower range, often under 20 µg/mL, for dilute samples. Run every standard and every unknown sample in at least duplicate, ideally triplicate — BCA color development has enough well-to-well variability that a single reading per point makes a bad well hard to catch.
2. Mix the working reagent
Combine Reagent A and Reagent B at the kit’s specified ratio (commonly 50:1) and mix thoroughly — the working reagent should be a clear apple-green color before it contacts any protein. Prepare only the volume needed for the plate; leftover mixed reagent degrades and should not be stored for a later run.
3. Add working reagent and incubate
Add working reagent to each standard and sample well (a common ratio is 8 parts reagent to 1 part sample), mix, and incubate. Two incubation regimes are in common use: 30–60 minutes at 37°C, or an accelerated ~30 minutes at 60°C for a faster, slightly less precise read. Whichever is used, hold every well — standards and samples — to the same time and temperature, since the purple color continues developing slowly even after the nominal endpoint.
4. Read absorbance at 562 nm
Cool the plate briefly to room temperature if it was heated, then read absorbance at 562 nm. Subtract the mean blank (diluent-only) absorbance from every standard and sample reading before proceeding to curve-fitting.
5. Build the standard curve and calculate concentration
Plot blank-corrected absorbance against known BSA concentration. Unlike Bradford, which is reasonably linear across its working range, the BCA standard curve is characteristically slightly non-linear, especially toward the top of the range — a second-order (quadratic) polynomial fit typically tracks a BCA curve better than a straight line, and most plate-reader software offers this as a built-in curve-fit option. Once the curve is fit, solve it for each sample’s blank-corrected absorbance to get a concentration, then multiply by any dilution factor applied before the assay to get the concentration in the original sample.
Common interferences and how they show up
- Reducing agents (DTT, β-mercaptoethanol, TCEP): these reduce Cu²⁺ independently of protein, inflating readings or preventing a usable standard curve. Some vendors sell a reducing-agent-compatible BCA reagent for exactly this case; otherwise, remove or dilute the reducing agent below its interference threshold, or switch to Bradford.
- Metal chelators (EDTA, EGTA): these sequester the copper the assay depends on, suppressing color development and understating concentration. As with reducing agents, a compatible reagent variant exists for some kits, or the chelator needs to be diluted out or removed.
- Detergents: BCA tolerates SDS and most common detergents far better than Bradford does, which is the main reason BCA is the default for lysates prepared in an SDS-containing buffer — but very high detergent concentrations can still shift results, so check kit-specific tolerance limits rather than assuming unlimited tolerance.
- Lipids and membrane components: BCA can respond to lipids and phospholipids at meaningfully high concentrations, which is worth flagging for membrane-fraction preps rather than clarified lysates.
Frequently asked questions
What does BCA stand for in a protein assay?
Bicinchoninic acid — the reagent that chelates the Cu¹⁺ produced when protein reduces Cu²⁺, forming the purple complex the assay measures at 562 nm.
Is BCA more accurate than Bradford?
Neither assay measures an absolute, protein-independent quantity — both report a result relative to whatever standard (almost always BSA) was used to build the curve. BCA’s color response tends to vary less from one protein to another than Bradford’s does, which is often described as better protein-to-protein uniformity rather than greater absolute accuracy. The better choice for a given sample usually comes down to compatibility with the buffer, not a general accuracy ranking.
Why is my BCA standard curve not linear?
Some curvature, especially at the higher end of the range, is expected and characteristic of the BCA chemistry — it is not automatically a failed run. Fit the curve with a quadratic (second-order polynomial) function rather than forcing a straight line, and treat only samples that fall within the standard curve’s tested range as reliable; extrapolating past the top standard is a common source of error.
Can I use BCA on a sample with SDS in the buffer?
Generally yes — BCA is substantially more detergent-tolerant than Bradford, which is why it is the common default for SDS-lysed samples. Check the specific kit’s documented detergent tolerance limit before assuming an unusually high SDS concentration is safe.
How long does the BCA working reagent stay usable once mixed?
Treat freshly mixed working reagent (Reagent A + Reagent B) as good for the working day it was prepared, not for storage — mix only the volume the plate needs and discard the rest rather than saving it for a later run.
Once total protein concentration is established, the next step for most workflows is normalizing loading before a Western blot — see Western blot loading controls for housekeeping-protein vs. total-protein normalization — or before setting up a sandwich ELISA. For cell-based samples, confirm viable cell counts with a hemocytometer count before lysis, so protein yield and cell number can be cross-checked against each other.
