FACS buffer is a simple solution built on phosphate-buffered saline (PBS) with a protein source and, depending on your application, a chelating agent and preservative. The standard version takes about 10 minutes to prepare. The exact recipe shifts slightly based on whether you’re staining for analysis, sorting live cells for culture, or fixing samples, so understanding each component lets you tailor the buffer to your experiment.
The Standard FACS Buffer Recipe
A widely used formulation for 1 liter of FACS buffer is:
- 1x PBS as the base, brought to 1 liter
- 2 g BSA (bovine serum albumin), giving a final concentration of 0.2% (w/v)
- 2 mL of 0.5 M EDTA, giving a final concentration of 1 mM
- 0.1% sodium azide (optional preservative)
Some protocols substitute FBS (fetal bovine serum) for BSA at 5–10% by volume. A Nature Protocols method, for example, uses 5% FBS plus 1% penicillin-streptomycin in PBS, filtered through a 0.22 µm filter and stored at 4 °C for up to two weeks. The choice between BSA and FBS matters, and we’ll cover that below.
What Each Component Does
PBS keeps the solution at a physiological pH (around 7.4) and osmolarity that won’t damage cells. It’s the inert backbone of the buffer, and you can use a commercial 1x solution or dissolve tablets or powder yourself.
The protein component, either BSA or FBS, serves two purposes. First, it blocks nonspecific antibody binding. Antibodies can stick to plastic tubes and cell surfaces in ways that create background noise. Protein in the buffer occupies those binding sites. Second, it cushions cells and helps maintain viability during the repeated wash and centrifugation steps of a staining protocol.
EDTA chelates (grabs) calcium and magnesium ions in solution. Without those ions, cells are far less likely to clump together. Clumps clog the flow cytometer’s nozzle and produce inaccurate scatter data. A concentration of 2–5 mM is typical for sorting applications.
Sodium azide at 0.1% prevents bacterial growth, extending the buffer’s shelf life at 4 °C. It also stops cells from internalizing surface-bound antibodies during staining, which keeps your signal on the cell surface where you want to measure it.
BSA vs. FBS: Which Protein to Use
BSA is a purified, defined protein. Because its composition is consistent from lot to lot, it produces lower and more predictable autofluorescence. That matters when you’re working with dim markers or need high resolution between populations. The University of Wisconsin flow cytometry lab recommends using a minimal amount of BSA (0.1–1%) specifically to keep autofluorescence low and improve population separation.
FBS is cheaper and contains a broader mix of proteins, growth factors, and nutrients. That broader mix is helpful when you want to keep cells happy for downstream culture. The tradeoff is lot-to-lot variability and slightly higher background fluorescence. Non-dialyzed FBS also reintroduces calcium and magnesium, which can promote cell clumping. If you use FBS for sorting, dialyzed FBS at 1–5% avoids that problem.
For routine analytical staining where cells will be fixed afterward, either works fine at the concentrations listed above. For sorting cells that need to stay alive and functional, BSA gives you more control over what’s in the tube.
Step-by-Step Preparation
Start with 900 mL of 1x PBS in a clean glass bottle. Add 2 g of BSA (or your chosen protein) and stir gently until fully dissolved. Vigorous stirring creates foam, which denatures protein at the air-liquid interface, so use a low-speed magnetic stir bar or swirl by hand.
Add 2 mL of 0.5 M EDTA stock solution. If you’re including sodium azide, add 1 g per liter (for a 0.1% final concentration). Top up to 1 liter with PBS and mix.
Filter the entire volume through a 0.22 µm vacuum or syringe filter. This removes particulates and sterilizes the buffer in one step. Particulates are a real problem on a cytometer: they show up as events in your data and can block fluidics lines. Store the finished buffer at 4 °C and use it cold. Cold buffer slows down cellular metabolic activity, reducing antibody internalization and keeping cells in better shape during staining.
When to Leave Out Sodium Azide
Sodium azide inhibits cellular metabolism by blocking an enzyme in the mitochondrial respiratory chain. That’s useful for staining, but it’s toxic to live cells you want to recover. If you’re sorting cells for culture, functional assays, or any downstream work that depends on cell health, leave sodium azide out entirely. Without it, the buffer has a shorter shelf life (roughly one to two weeks at 4 °C), so make smaller batches and watch for cloudiness, which signals contamination.
Handling Sticky or Clumpy Samples
Dead and dying cells release DNA into solution. Free DNA is extremely sticky and causes surviving cells to clump into aggregates that will clog a sorter nozzle and ruin your data. This is especially common with freshly isolated primary cells from tissues.
The fix is adding DNase I to your buffer. For routine prevention, add 25–50 µg/mL of DNase I directly to your FACS buffer. DNase requires magnesium to work, so include 5 mM magnesium chloride as well. Note that if you’re relying on EDTA to prevent clumping, you have a conflict: EDTA chelates the magnesium that DNase needs. In samples with significant cell death, the better strategy is to drop EDTA from the buffer and rely on DNase plus magnesium instead.
For severely sticky samples, a pre-treatment step helps. Incubate cells in 100 µg/mL DNase I (or 10 units/mL) with 5 mM magnesium chloride in balanced salt solution for 15–30 minutes at room temperature. Then resuspend in your working FACS buffer containing at least 1 mM magnesium chloride and 25–50 µg/mL DNase to keep aggregation under control throughout the staining and sorting process.
Adapting the Buffer for Sorting
Sorting puts cells under more physical stress than analysis. They pass through a narrow nozzle at high pressure, get charged with an electrical field, and are deflected into collection tubes. Your buffer needs to support viability through all of that.
Keep protein concentration on the lower end (0.1–1% BSA or 1–5% dialyzed FBS) to minimize autofluorescence while still cushioning cells. Use EDTA at 2–5 mM to prevent adhesion, unless your sample has high cell death, in which case switch to the DNase strategy described above. Leave out sodium azide. And filter everything: buffer, collection media, and any additives. A single fiber or crystal of undissolved salt can block the sort nozzle and shut down your experiment.
For the collection tube, fill it with a richer medium. Sorted cells land in a small volume at high velocity, and a cushion of culture medium with 10–20% FBS absorbs that impact and gives cells nutrients to start recovering immediately.
Storage and Shelf Life
FACS buffer with sodium azide stays usable at 4 °C for several weeks. Without azide, plan on one to two weeks maximum. Always store it cold, and bring it to the bench cold on the day of your experiment. If you see any cloudiness or floating particles, discard the batch and make fresh buffer. Protein-containing solutions are a good growth medium for bacteria, and contamination introduces particles that mimic cells on the cytometer.
For labs that use FACS buffer infrequently, making 100–200 mL at a time avoids waste. The preparation is quick enough that making it fresh every week or two is practical.