When working with patients, samples are rarely collected from all of the patients on the same day. That leaves two options for running samples: cells can be collected and run on the same day over the course of the study or cells can be cryopreserved and run at one time. For cryopreserved blood samples, peripheral blood mononuclear cells (PBMCs) are usually the preferred method. However, we’ve found an alternative way to preserve whole blood using the Proteomic Stabilization Buffer by Smart Tube. This buffer was originally created for studying cell signaling using mass cytometry. When we tried out this buffer on cells analyzed by flow cytometry, we were quite impressed at its ability to preserve the forward and side scatter properties of the populations (including neutrophils!) after thawing (Figure 1). In fact, we’ve started to refer to it as “magic goo”. (Note: this buffer does fix the cells prior to freezing.)
Using the “magic goo” is relatively straightforward. To preserve the blood, 1 mL of blood is mixed with 1.4 mL of Proteomic Stabilization Buffer, incubated for 10 minutes, and frozen. Then when the samples are ready to be used, they are thawed and washed with the provided thaw-lyse buffer. Smart Tube does have optimized Smart Tubes containing 1.4 mL of “magic goo” and a base station for incubation steps. These are a great option for ensuring that samples are processed the same way every time or for anyone who needs to stimulate their cells prior to freezing. If you don’t want to use their Smart Tubes, they also sell the “magic goo” in bulk.
I did run into a few small issues that took me a while to sort out. First of all, I discovered that the thawing time was absolutely critical in the success of the samples. Smart Tube does note in their protocol that it is important to move to the next step after the samples have been thawed. What they don’t specify is that if too much time passes after the samples are fully thawed, the sample will turn black and clump, making it impossible to pipette. I learned that lesson pretty quickly! Now I make sure to add the thaw-lyse buffer immediately after the samples have thawed to avoid the clumping problem.
The main problem that I encountered was a tough one that stumped me for quite a while. As I was titrating antibodies, I kept finding high levels of nonspecific background staining in all cell types. It seemed to occur with only some of the antibodies, but I could not figure out why it happened or how to prevent it. Finally, a Fluidigm Application Scientist suggested I try using heparin during my blocking step and it worked fantastically! I followed a paper by Rahman et al. which describes using heparin to block charge-based interactions and reduce background staining on eosinophils. The figure below shows my initial test for adding heparin in the blocking step and increasing the FBS in the staining buffer (Figure 2). As you can see, the CD4 antibody was staining all of the cells in the blood, but the addition of heparin decreased the background in every cell type while maintaining the fluorescence intensity of the positive population.
Since learning about this heparin blocking method, I’ve heard that some of the researchers using our core have tested it out on their own experiments and it appears to have broad applications. If you’re interested in trying it out, here’s the protocol I used:
- Block cells in Fc block and 200 U/mL heparin in a total volume of 50 μL FACS staining buffer
- Incubate 20 minutes at 4°C
- Add 50 μL of antibody master mix for a final staining volume of 100 μL
- Incubate 30 minutes
- Wash and run samples
Let us know if you’re interested in using any of the Smart Tube products! The CAT Facility has the Smart Tubes and base station. We would also love to hear if you have tried out heparin during your blocking step and what applications it is useful for!