This guide explains how to use the Protein assay & loading calculator: it turns BCA or Bradford plate readings into lysate concentrations and SDS-PAGE loading volumes. It also shows exactly how every number is calculated.
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What you need
- Absorbance readings for your standard series and samples, usually exported from the plate reader to Excel.
- The dilution you used for the assay, if you diluted the lysate before measuring it.
- How much protein (µg) and what volume (µL) you want in each lane.
How it works
- Paste the plate. Copy the block of absorbance values from Excel or your reader export. Click the blue box and paste with Ctrl+V (⌘V on a Mac). You can copy the row letters (A–H) and column numbers (1–12) too. The calculator uses them to put each value in the right well. 96- and 384-well layouts both work. If your export holds several reads, copy only the one you want, e.g. the 562 nm read for BCA.
- Mark the standards. Drag across the standard wells, or click the first well and then the last.
- The shape of the block sets the replicate layout. A block two columns wide and eight rows tall is read as duplicates side by side, one standard per row.
- Standards are labelled S1, S2… on the plate, so you can check the order at a glance.
- Check the concentrations. A default series is filled in automatically.
- With 9 standards the defaults are 0, 25, 125, 250, 500, 750, 1000, 1500 and 2000 µg/mL. With 8 standards the series is the same without 25.
- If S1 reads darker than the last standard, the series is filled in highest first.
- Type over any value, or press Reverse order, if your series is different.
- Mark the samples. Drag across the sample wells, and drag again to add another block. Samples are numbered 1, 2, 3… following the replicate direction:
- down each pair of columns when replicates sit side by side;
- across each pair of rows when replicates sit top to bottom.
- Check the curve. Choose BCA or Bradford and a curve fit. The equation, R² and the blank value appear under the chart, and your samples are plotted on the curve as orange diamonds.
- Set the loading mix. Enter the assay dilution, the µg and µL per lane, and how many times the lane mix to make. The table shows two sets of volumes side by side:
- per-lane volumes, for loading a single lane;
- whole-tube volumes, for preparing a stock you can load several times.
- Export. Export CSV saves the raw plate, the wells used, means, CVs, concentrations and every volume, ready to paste into your lab notebook or Excel. Print / PDF prints just the calculator.
Marking wells
- Exclude. Click a well to leave it out of the mean, for example one with an air bubble or a pipetting error. Excluded wells are struck through on the plate and listed in the CSV. Click again to bring a well back.
- Edit values. Type absorbances straight into the wells, or paste a block of cells into any well. The block fills across and down from that well. Use this if you read samples one at a time on a spectrophotometer.
- Replicates. The replicate count (1–3) and direction are set from the standards block. Change them in the toolbar if they were guessed wrong.
- Reusing a layout. Pasting a new plate keeps the standard and sample wells you marked. If you use the same layout every time, you only mark the wells once per session.
The calculation
Blank-corrected A = mean A − mean A of the 0 µg/mL standard
Assay concentration = read off the standard curve (µg/mL or mg/mL)
Lysate concentration = assay concentration × assay dilution (µg/µL)
(÷ 1000 first if the standards are in µg/mL; 1 mg/mL = 1 µg/µL)
Per lane
Lysate = µg per lane ÷ lysate concentration
5× sample buffer = µL per lane ÷ 5
Lysis buffer = µL per lane − 5× sample buffer − lysate
Whole tube = each per-lane volume × scale-upCurve fits
- Linear: A = a + b·x, solved for x.
- Quadratic: A = a + b·x + c·x², solved on the rising part of the curve.
BCA responds almost linearly over the usual 0–2000 µg/mL range, so linear is the default. Bradford curves flatten at higher concentrations, so the default for Bradford is quadratic. If the points sit clearly off the line, switch fits. If a quadratic curve turns over inside your standard range, the calculator flags it. In that case use a linear fit or drop the top standard.
Replicates
Replicate wells are averaged, and the coefficient of variation (CV) is shown next to each mean. A CV above 10% is highlighted. It usually points to a pipetting error or a bubble, and excluding that one well is often the fix.
Why the 5× buffer is always one fifth
The sample buffer has to end up at 1× in every lane. So 5× buffer is always one fifth of the final volume, however concentrated the sample is. Lysis buffer makes up the rest, so every lane has the same volume and the same buffer composition. The calculator assumes 5× sample buffer.
When a sample is too dilute
After the sample buffer, only so much room is left for lysate:
Maximum µg per lane = lysate concentration × (µL per lane − µL per lane ÷ 5)For 30 µg in 10 µL, 2 µL goes to 5× buffer and 8 µL is left for lysate. The lysate therefore needs at least 30 ÷ 8 = 3.75 µg/µL. A sample below that is marked too dilute. Its lysate and lysis buffer volumes show “—”, and the message under the table gives the most it can carry. You have three options:
- load fewer µg in every lane;
- load a larger volume per lane, if your wells allow it;
- concentrate the lysate.
Keep the µg the same in every lane you plan to compare.
Assay dilution
Diluting the lysate before the assay keeps readings inside the standard range. It also dilutes substances that interfere with the assay. Enter the dilution factor, not the ratio of lysate to diluent. For example, 5 µL lysate + 45 µL diluent is a 10× dilution. The factor applies to every sample, and you can change it for a single sample in the table’s Dil.× column.
Warnings
| Message | What it means | What to do |
|---|---|---|
| too dilute | The requested µg does not fit in the lane volume. | Load fewer µg, use a larger volume, or concentrate the lysate. |
| above top std | The reading is above the highest standard, so its concentration is extrapolated. | Dilute the lysate further and re-assay. |
| below lowest std | The reading is below the lowest non-zero standard. | Use less dilution, or add a lower standard. |
| ≤ blank | The sample reads at or below the 0 µg/mL standard. | Check the well and the dilution. |
| CV highlighted | The replicates disagree by more than 10%. | Look for a bubble or pipetting error, and exclude that well. |
| R² low | The standards fit the curve poorly (R² < 0.98). | Check the concentrations and their order, try the other fit, or exclude an outlier. |
| Under 1 µL per tube | A volume is too small to pipette accurately. | Raise the scale-up, or pre-dilute the lysate. |
Frequently asked questions
Is my data uploaded anywhere?
No. The calculator runs entirely in your browser, and nothing is sent to a server or stored. Closing the tab ends the session, so export the CSV before you leave.
Which plate readers does it work with?
Any reader whose results you can copy into Excel or a text file. The calculator reads tab-, comma- and semicolon-separated values, with or without row letters and column numbers. Cells with text such as OVRFLW are kept on the plate but ignored in the calculation.
My standards run across the rows, not down the columns.
That works too. Drag across them, and a block two rows tall is read as duplicates stacked top to bottom, one standard per column. Samples then follow the same direction.
Can I use a different standard series, or a protein other than BSA?
Yes. Type or paste your own concentrations into the standards table, and pick µg/mL or mg/mL to match them. If the numbers look like the other unit (for example 0–2 when µg/mL is selected), the calculator asks you to check before you go on.
BCA or Bradford — which should I use?
It depends mostly on what is in your lysis buffer:
- BCA tolerates most detergents but is affected by reducing agents such as DTT and β-mercaptoethanol, and by chelators such as EDTA.
- Bradford is affected by detergents such as SDS.
Either way, measure the lysate before you add sample buffer, which contains both SDS and a reducing agent.
Why don’t I need to name my samples?
Each sample is identified by its number, which matches the number shown on its wells on the plate. The CSV lists the wells for each number, so you can add names in Excel afterwards.
How to cite this tool
If you use these values in a publication, you can describe the method like this:
Protein concentrations were determined by BCA assay against a BSA standard curve, and loading volumes were calculated with the PYMtools Protein assay & loading calculator (https://pymtools.castingfilter.com/protein-assay-calculator/, accessed DD Month YYYY).
Related tools
- Western blot quantification: background-corrected band intensities, normalised to your housekeeping protein
- ΔΔCt calculator: relative qPCR quantification (coming soon)
For research use only. Not for use in diagnostic procedures. Results are provided without warranty — verify them before publication.