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6 Tips for Improving Sample Staining for Flow Cytometry
How To Guide

6 Tips for Improving Sample Staining for Flow Cytometry

6 Tips for Improving Sample Staining for Flow Cytometry
How To Guide

6 Tips for Improving Sample Staining for Flow Cytometry

Flow cytometry is a single cell technology that utilizes lasers and fluorescence (in most cases) to study the gene expression profile and function of cells. The power of flow cytometry is its ability to collect data at a high rate (up to 30,000 cells per second) and in multi parameter (commonly 15 or 16 but extending to 40+ in a single sample). Developed initially in the field of immunology, flow cytometry now has applications throughout the biological sciences and beyond. A successful flow cytometry experiment begins with good sample preparation.

1. Source of sample


Flow cytometry measures the characteristics of individual cells as they pass through a series of lasers one at a time. This requires samples to be in a single cell suspension. If we are studying cells from the blood or perfusate then generating a single cell suspension is relatively straightforward as the cells are inherently not physically attached to one another. However, biologists are often interested in the cells associated with tissues and this makes generating a single cell suspension more of a challenge.

Tissue is a collection of different cells intimately linked together in a complex. We need to tease this complex apart whilst maintaining cell viability, physiological homeostasis and antigen integrity to be able to perform accurate flow cytometric analysis. Some tissue, such as spleen for example, is relatively soft and is easily disaggregated by pushing it through a 100µm mesh with the back of a syringe plunger or similar. As this involves no enzymic actions cell viability is maintained and antigens are unaffected.

The majority of tissue however is considerably more challenging! In these cases, a bit more ‘muscle’ is required to disaggregate the cells and enzymes, temperature and mechanical techniques are commonly used, often together, to produce single cells. These approaches can be very harmful to the cells and result in extensive cell death and loss of antigen.

Protocols for performing tissue digestion are available online from a number of sources. However, a highly comprehensive guide to all things tissue digestion can be found on the Worthington Biochemical Corporation website.1 This is a great resource for initial guidance to disaggregating a wide variety of tissue from a long list of organisms.

The general method for digesting tissue follows a protocol that involves mechanical ‘mincing’ of the tissue, followed by enzymic digestion and finally filtering.

To understand the protocol, we need to consider the potential pitfalls of each step.

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2. Mechanical techniques


After excising the tissue from the organism, dissect away as much of the unwanted fat and tissue as possible to improve the effectiveness of the rest of the protocol. Mechanical techniques such as mincing with a scalpel or chopping with scissors followed by syringing and filtering are used to coarsely break up the tissue and increase the surface area accessible to enzymes. It is important that mechanical disruption is done in the presence of a buffered medium suitable for the enzymic stage of the protocol. The enzyme may, for example, need divalent cations to function, in this case you should avoid chelators such as EDTA. If trypsin is used, you should use a balanced salt solution that does not contain magnesium or calcium. Directions for the most suitable buffer should be provided with the product. If not, suitable buffering information can be found with a little research.

3. Enzymic digestion and incubation temperature


These two techniques are discussed together because enzymes are active at physiological temperatures, which is where cells are often at their most active. There is therefore a potential for incubation periods to increase sample death as cells are out of their physiological environment and may become stressed causing aberrant signaling or changes in their physiology (receptor internalization or activation). Commonly, collagenase type I is used for digesting tissues, but other enzymes such as trypsin or papain can be used. All three are examples of proteolytic enzymes which cleave proteins at particular sequences. Be careful, if the cleavage sequence is present in your target antigen, it too will be cleaved and therefore absent from downstream analysis.

From the physiological aspect, it is important that incubation temperatures and times are optimized to enable the enzymes to do their ‘thing’ without adversely affecting the sample. It’s difficult to say how long samples should be incubated to obtain optimal digestion, commonly 1 or 2 hours at 37°C is sufficient. Incubation should allow enough time for the tissue to be digested but not so long that cell death increases - this will vary depending on the type of tissue. Keep a close eye on the process and stop the digestion once your sample appears homogeneous as over digestion is never a good idea. Be aware, incubation periods of up to 18 hours are necessary for some types of collagenase. 

Optimizing incubation times and temperatures is often an empirical process that requires a careful balancing act and an element of trial and error. Cell degradation may be reduced by reducing incubation temperature but consequently this may reduce enzyme activity. Increased incubation time would then be required which may then once again increase cell degradation.

Trypsin is a strong tissue disaggregating enzyme and can result in loss of the sample if incubated for too long at 37°C. With trypsin it is recommended that you perform an overnight incubation at 4°C to enable the enzyme to penetrate the tissue. The sample should then be incubated for 30 mins at 37°C the following day to break it down.

A word on enzymes - and what to consider. Enzymes are biological agents and as such will contain variations in activity. The quality of the enzyme can vary between manufacturers with regards to how pure they are. Enzymes can also vary from lot to lot, which can be frustrating when working with them over a long period of time. When purchasing your enzymes ensure you get sufficient to perform both the optimizations and as many experiments as you can possibly imagine you’ll need! Alternatively, contact the manufacturer as many are accommodating in holding stock of specific lots on request.

Following the digestion step we quench the activity of the enzyme by adding balanced salt solution (e.g. Hanks Balanced Salt Solution) with approximately 2% serum (e.g. fetal bovine serum, (FBS)) to the sample. You should at this point have a loose tissue preparation that can be separated into single cells by gentle mixing with a 1mL or 10mL pipette (depending on your volume).

Recover the cells from the initial suspension by centrifugation at about 300-400 x g for 5 mins (with the brake on) to remove the residual enzyme from the cells. After resuspending the resultant pellet in balanced salt solution pass the sample through a suitable filter (we often use a 100um filter) to remove any final aggregates from the sample and perform a cell count/ viability assessment (see below). Ideally all steps after the digestion would be performed on ice or at reduced temperatures with chilled buffers and a cold centrifuge to avoid any further cellular activity and minimize cell death.

4. Dead cells


Dead cells can be a real problem in analysis and can be a major issue when it comes to good staining with antibodies for flow cytometry.

Dead cells readily bind antibodies non-specifically generating artifactual data and false positive staining. They also, in the process of dying, release large amounts of DNA into the culture – giving the sample a think “snotty” appearance – which can cause cells to stick together and in turn die. If this occurs, liberated DNA can be removed easily using DNase which can be added to many tissue preparation buffers.

Following the digestion protocol it is a good idea to check the cell viability. This can be done with a number of quick and simple methods. Traditionally Trypan Blue is gently mixed with an aliquot of the cell sample prior to its application to a hemocytometer (counting chamber slide) and acts as a differential stain to assess the viability of the sample. When viewed down a light microscope Trypan Blue highlights dead cells with an intense blue color and allows the counting of non-blue (live) and blue (dead) cells. Recently highly efficient systems based on fluorescence discrimination of live and dead cells have become available which employ a built in microscope and camera to do the counting and assess viability. A stain such as acridine orange is used to label all the cells and propidium iodide or DAPI can be used to stain the dead cells. Employing a camera and software makes the counting robust and much more straightforward than a hemocytometer.

If dead cells are a major issue in your sample, there are commercially available enrichment kits which can be used to "clean up" the sample in a similar way to antibody-based sorting using beads. These kits work well, dramatically improving sample staining and by minimizing the presence of undesirable dead cells reduce acquisition time and file size.

A number of companies now sell fixable viability reagents. Historically if you wanted to ensure you were not looking at dead cells in your analysis you would add a dead cell discriminator such as propidium iodide or 7AAD. These reagents however only worked on live samples – after a long day isolating cells and staining samples the last thing you want to do is run the samples at midnight! Fixable viability reagents are great as they can be used prior to staining and then fixed. Samples can then be stored overnight and run the next day when there may be more time available on the machine and you have more energy! Being able to fix samples also makes them safer to handle from a biosafety perspective.

5. Red blood cells and debris


Red blood cells (RBCs) are abundant in many biological systems and can be problematic from the point of view of sample acquisition. RBCs outnumber non-RBCs to a ratio of ~1000:1 in most sample types, and can mask events that we would be interested in. They can be relatively simple to remove with an osmotic lysis step, but beware – some lysis buffers can be harmful to non-RBCs and may result in loss of cell types such as mast cells or basophils.

Debris, generated by disaggregating tissue and from the breakdown of dead cells during sample prep, can also be a problem when analyzing tissue. In a similar way to RBCs, large amounts of debris can make identification of cells a challenge. Debris is a little more difficult to remove but can be done by density gradient centrifugation or using sorting techniques such as enrichment beads.

6. Getting the best from your flow cytometer


An important aspect of flow cytometry sample preparation that is often overlooked is optimization of the staining conditions. It is important to use the correct amount of antibody for the sample. Perform titrations of all your antibodies – this can seem like an expensive exercise but ultimately may save money as you may require less antibody than the amount per test suggested by the manufacturer. Often companies validate antibodies against samples which potentially give the best results – this is often peripheral blood mononuclear cell (PBMC) samples. This doesn’t necessarily mean that this is the best concentration for your tissue of interest. 

Count your cells. The number of cells being stained has a major impact on the quality of the staining. Where possible ensure you always label the same number of cells – this isn’t always easy. If you are working with small biopsies or bodily fluid the number of cells can vary dramatically, in these situations it’s a case of optimizing to the likely highest number of cells you would expect to have in a sample. 

In summary, the best way to improve sample staining for flow cytometry is to ensure you have a healthy, single cell suspension that has minimal levels of dead cells and is free from RBCs and debris as much as possible. Use a viability reagent so that you can identify dead cells in your sample and remove them from the analysis pipeline to minimize artefacts from non-specific antibody binding and aggregates. Finally, take the time to perform titrations of your reagents to set numbers of cells, this will save you time and money in the long run and improve the quality and allow standardization of the data you generate.

References

1. Worthington Biochemical Corporation web site http://www.worthington-biochem.com
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