How To Prepare a Good Single Cell Suspension

blood being placed in centrifuge to symbolise single cell suspension

Table of Contents


Flow cytometry is a powerful technology that encompasses many applications and techniques. For all these applications, cell processing techniques are required to produce a cell suspension from a diverse range of cell types or tissues. Producing a good single cell suspension is needed to achieve high quality flow cytometric analyses. Typically, once a single cell suspension has been produced, further cell treatment and staining is required prior to running the samples on the flow cytometer. This blog will describe how to prepare cell suspensions using examples from different starting biological materials. I will outline some useful methodology, hints and tips on how to analyse the quality of the cell suspension and I will also make recommendations for troubleshooting the preparation of difficult samples.

Why it is important to prepare a good single cell suspension

There are several important components of a successful flow cytometry experiment. These can be divided into good sample preparation, instrument setup and maintenance, data acquisition and analysis. The first step in preparing a good sample is the preparation of a good single cell suspension, this is frequently followed by various staining techniques.

What are the reasons why a good single cell suspension is needed for high quality flow cytometry?

1.1 To minimise clumps which can cause instrument blockages. Clumps can stop the sample running or prevent accurate positioning of cells within the sensing area causing inaccurate scatter and fluorescence measurements.

Figure 1 1
Fig 1.
Schematic of a typical scatter plots of a homogeneous sample showing a cell line. Forward scatter (FSC) and side scatter (SSC) are displayed on linear scales. (A) is a sample run with no blockage in the flow cell, (B) is the same sample run with a partial blockage in the flow cell. Healthy cells are within the set gate in (A) and dead and unhealthy cells have lower FSC than the healthy cells. In (B) the healthy cells from the same sample shows lower FSC and SSC because the flow cell has become partially blocked. 

1.2 To achieve uniform staining and fixation. When clumping occurs, unless the clumps are dispersed prior to staining and fixation, access of reagents to individual cells may be impaired resulting in reduced staining of the cells within the clumps. This may in turn cause uneven staining or fixation of the cell sample. If the clumps are subsequently dispersed prior to running the sample, the uneven staining or fixation can be seen.

Figure 2 2
Fig 2: diagram showing the effects of uneven fluorescence staining resulting in bimodal fluorescence intensity.

1.3 To ensure there are sufficient single cells in each cell sample for a statistically significant analysis. The cell numbers required for statistical significance will depend on several factors: the percentages of cells to be analysed, the type of analysis and the variability of the measurement. For example, more cells are required for rare cell analysis or for the analysis where there is only a small shift in fluorescence such as phospho-protein analysis.

1.4 To avoid preferential loss of some cell types within samples made up of a mixed population. This may include more fragile and adherent cell types that are more prone to clumping eg. cell samples containing dendritic cells, neutrophils, monocytes and macrophages. A knowledge and understanding of these cell characteristics can help to choose the most appropriate techniques to prepare a good single cell suspension. For example, the analysis of whole blood rather than purified subpopulations from blood can minimize the loss of rare and fragile cell types such as dendritic cells. In addition, less sample manipulation can reduce cell activation of easily activated cell types such as neutrophils and the proteins present in whole blood can help to maintain good cell viability.

Figure 3AB
Fig 3.
Schematic diagram (A) of a density plot of forward scatter (FSC) vs Side scatter (SSC), set on linear scales. The populations typically present in a lysed human whole blood sample. The granulocytes population includes neutrophils and eosinophils. These cell types are prone to activation and cell death during rigorous sample processing leading to changes in scatter and cell loss. Monocytes can adhere to plastic of the sample tube or well and so resuspension prior to the analysis is important to avoid cells lost from the analysis. Therefore, the preparation of samples requires careful cell handling to minimize changes in the analysis including cell scatter and cell loss.
NB. the relevant positions can change slightly between instrument types and lysis method used. The real data shown in (B) shows these typical scatter differences as seen in the schematic (A).

Description of the main categories of cell suspensions

There are 3 main categories of cell suspensions prepared for flow cytometry analysis:

  • Dispersal of cells from tissues
  • Cell suspensions from cultured adherent cells
  • Cells prepared from pre-existing cell suspensions

For each of these categories and for particular cell types within each category, there are different considerations for achieving a good single cell suspension. These considerations and recommendations will be discussed with these cell categories in mind.

Recommendations for each category

2.1 Dispersal of cells from tissues: The techniques used involve either mechanical or enzymic tissue disaggregation or a combination of both. The ease of cell dispersal and isolation varies depending on the tissue type. Various published methodologies are available for particular tissues and cell types. I have used and optimized a range of these methods for preparing a variety of cell suspensions.

These include:

  • The gentleMACS™ “Tubes” and “Dissociator” are a benchtop system for the semi-automated dissociation of tissues into single-cell suspensions. Pre-installed protocols are provided for particular tissues and cell types. Further optimisation can be achieved by modifying the time for the dissociation.

Images from Miltenyi Biotec.

Fig 4. Shows a GentleMACS instrument from Miltenyi Biotec on the left and specialised tubes for cell dissociation and a petri dish has a fragment of tissue to be dissociated in the tubes.

2.2 Cell suspensions from cultured adherent cells:

Cell detachments methods include:

  • Enzymic detachment: This is where enzyme-based solutions are used to detach adherent cells. An example commonly used for flow cytometry is Accutase or TrypLE.  These are usually safer to use than Trypsin, as unlike Accutase solutions, trypsin often cleaves peptides and degrades many cell surface proteins.
    • Non-enzymic cell detachment: These include mechanical cell detachment by scraping or the use of the cation chelator, EDTA solution or commercial cation chelator solutions such as Sigma dissociation buffer (Merck catalogue C5789).  

Abhilok Garget et al compared enzymic and non-enzymic detachment of human bone marrow–derived mesenchymal stromal cells (MSC) for various cell functions. For non-enzymic methods, there was better viability and surface receptors functions with cell dissociation buffer than EDTA solution or scraping. For enzymic methods, TrypLE stripped chemokine receptors off the cell surface but Accutase did not.

Figure 5 1
Figure 5. Overlay histograms of adherent cells detached with trypsin or Accutase and immunofluorescently labelled with an antibody conjugate for one of the many cell surface antigens susceptible to degradation by trypsin. Time dependant degradation by trypsin is shown by reduced fluorescence staining intensity. However, when Accutase is used, fluorescence staining is not reduced.

2.3 Cells prepared from pre-existing cell suspensions

Pre-existing cell suspensions for flow cytometry include cell mixtures such as blood and bone marrow or cell lines grown as cell suspensions in culture. Cell suspensions may also be prepared by isolating cell populations, for example peripheral blood mononuclear cells (PBMCs) or lymphocyte subsets from blood.

Note, cell populations such as monocytes that exist as cell suspensions in blood can become adherent following incubation on polystyrene culture plates and differentiation of monocytes to macrophages increases adherence. Detachment may require rigorous pipetting for monocytes attached to plates or enzymic detachment for macrophages.

The preparation of cell suspensions for flow cytometry such as PBMCs or whole blood with or without red blood cell lysis, involves commonly used protocols. These are topics for other discussions and blogs.

How to prepare a good cell suspension-practical hints and tips

For all cell suspensions prepared from tissues, adherent cells and pre-existing cell suspensions:

3.1 How can viability be improved?

3.1.1 Add protein: A lack of protein in buffers or media during cell preparation frequently reduces viability, where some cell types are more susceptible than others. To achieve and maintain good viability, addition of protein is required at all stages of cell processing e.g. in wash and resuspension buffers or media. Typically, solutions containing 2% FBS or 1% BSA is suitable for immunofluorescence staining. Human AB serum may be preferable for applications where lower background activation of human primary cell cultures is needed. Exceptions to the addition of protein include cell preparation where protein can inhibit a process, such as when staining with fixable dyes is required for dead cell elimination from the flow cytometry analysis. Any protein present during the staining step can absorb dye limiting the amount available to stain the cells. Therefore protein should be added soon after this staining step.

3.1.2 Gentle resuspension of fragile cells: this is essential during sample preparation whilst still ensuring thorough mixing.

3.2 Which is the best way to remove clumps?

3.2.1 Add DNase: Damaged and dead cells release DNA. The DNA then acts like a glue causing aggregation of both dead and live cells, resulting in clumping. This reduces the number of single live cells in the sample for the analysis. Adding DNase breaks down the free DNA, thus reducing clumping and increasing the number of single cells available for the analysis. In addition, the use of DNase reduces the chance of partially or fully blocking the flow cytometer flow cell or sampling system.  A protocol I typically use is DNase (from bovine pancreatic DNase 1) at 25mg/ml final concentration added to the medium for cell isolation and final cell resuspension buffers. This can also be used in combination with staining techniques such as immunofluorescence staining.

3.2.2 Add EDTA: Cell adhesion in some cell types is cation dependent. When this occurs, clumping may also be a consequence. EDTA, a cation chelator can reduce this type of clumping. For these cell types, I typically add 2mM EDTA to buffers and media during cell preparations and staining. Exceptions should be made where EDTA interferes with cation dependent function such as integrin binding.

3.2.3 Cell mixing & pipetting for good single cell suspensions: Sufficient mixing is required at all stages of cell preparation, so it’s best to use gentle mixing of fragile cell types and don’t vortex at maximum speed. Cell mixing is more effective in smaller volumes than larger volumes, therefore it is better to vortex a cell pellet before the addition of buffer or media. For protocols that require fixation, thorough mixing is needed to obtain a good single cell suspension prior to the addition of fixative. Please note, steady addition of fixative and immediate mixing is required to reduce cell aggregation during fixation. Unless this is done, fixation can be uneven, and result in artifacts in the analysis of scatter or fluorescent staining. Washing to remove the fixative to prevent prolonged incubations in fixatives is often required. There are various fixation techniques that are used for different flow cytometry applications.

3.2.4 Mixing immediately prior to running samples on the flow cytometer: this is important to resuspend cells that have settled and may also be important to disperse recently formed clumps. This can be achieved by vortexing, flicking tubes rigorously or pipetting using 200μl tips just before loading the sample onto the cytometer manual sampling stage or using automated mixing features within microtiter plate loaders or tube racks.

Figure 6
Fig 6. Schematic shows the effects of even fixation of a good single cell suspension vs uneven fixation of a poor single cell suspension with clumps. (A) shows uniform fixation with homogeneous scatter, (B) shows uneven fixation resulting in bimodal scatter.

3.2.5 Using cell strainers: Some cell clumps may be too difficult to disperse using the methods already described. If so, then these clumps may need to be removed using cell strainers. This may be required at various stages including prior to cell isolation and staining as well as prior to running samples on the flow cytometer. If clumps are present prior to staining, removal of clumps produces more even staining. If clumps are present prior to running samples, removal of clumps reduces the chances of instrument blockages. The cell strainer can be chosen according to the pore size of its mesh and to accommodate the required sample volume. 70μl is a typical choice and various volume sizes of cell strainers are shown in Fig 7.

Figure 7
Fig 7. Cell strainers from various suppliers-Image from ThermoFisher

3.3 Polypropylene tubes and plates to reduce adherence: Where cell suspensions are prepared from adherent cells, cell adhesion to polystyrene plates and tubes may reduce the number of cells in suspension. This may occur prior to data acquisition where cells settle and attach to the plastic. If this occurs, adherence can be reduced by using tubes and plates made of polypropylene instead of polystyrene.

How to check the quality of the cell suspension prior to and during processing

It is advisable to check the quality of the cell suspension at different stages of the cell preparation, staining and sample acquisition.

4.1 Clumps can sometimes be large enough to be seen by eye: Initial checks of the quality of a single cell suspension can be done by looking at the appearance of the cell suspension in the tube or plate. This can be useful in assessing how successful the dispersal techniques or removal of large clumps has been e.g. before or after pipetting up and down, the use of DNase, EDTA or filtering with cell strainers. Large clumps are often visible by eye and this simple observation can frequently help to prevent blockages in the flow cytometer.

4.2 Use microscopy to check the cell suspensions: Simply using a low power light microscope to examine the cell sample and quality of the cell suspensions prior to and during processing can help to determine the best techniques required to produce a good single cell suspension. Monitoring cell health of cell cultures using low power light microscopy is often invaluable. Each adherent cell culture has a characteristic shape and appearance but any dead and dying cells float and clump together. The cell health of suspension cell cultures cannot be distinguished by the floating cells as all the cells float. However, the healthy suspension cells can be distinguished from the dead cells because they have a shiny appearance due to refraction of the microscope light source whereas the dead cell membranes appear ragged and darker due to less light refraction.

4.3 Analysis of the quality of the cell suspension by flow cytometry: Scatter measurements can tell you a lot about the condition of the cells, such as the relative proportion of clumps and debris in the cell suspension.

4.3.1 Analysis of scatter to access the quality of the cell suspension: Forward scatter is an indication of the size, shape and refractive index of the cells or particles, side scatter is a measure of the granularity of the cells or particles.

Figure 8
Fig.8 Schematic of forward scatter and side scatter measurements of a sample containing a single cell type. This alone can provide a lot of information about the amount of healthy single cells, clumps, unhealthy cells and debris that make up the cell suspension being analysed. Where the sample contains a mixed population of different cell types, using FSC and SSC alone to distinguish cell aggregates from larger cells and dead cells from smaller cells is more difficult.

Example of sample of a mixed population

Figure 9
Fig 9.
Peripheral blood mononuclear cells (PBMCs) prepared from human peripheral blood showing a typical scatter plot of (FSC) vs Side scatter (SSC), both set on linear scales. Note: It is important to identify the scatter of the cells of interest. In this example the gate has been set around the lymphocytes. A large gate has been deliberately set to include both activated (lymphocyte blasts) as well as the non-activated (resting lymphocytes). If the flow cytometry operator is not familiar with the scatter characteristics of the different cell types, important subpopulations may be lost in incorrect gating strategy. For example, larger cell types may be confused with cell aggregates and clumps.

Use of back-gating using a lymphocyte marker staining such as CD3 is a useful technique to ensure the correct scatter gate has been set.

Screenshot 241
Fig 10. shows back-gating of human blood using Lymphocytes (green) and monocytes (blue) and granulocytes (red) gates in CD3 vs CD14 plot displayed in the FSC vs SSC scatter plot. This allows the scatter gating strategy to be confirmed. Plots from Bio-Rad.

In addition to using FSC vs SSC to analyse aggregated and clumps, single cells and doublets can be distinguished using a combination of height and area or height and width of the forward or side scatter signals. The theory is illustrated below.

Figure 11a
Fig 11a. Schematic showing how the FSC and SSC signals are produced for single cells and doublets. The area and width for the doublets are higher than for the singlets but the height for singlets and doublets remain the same. The resulting scatter plots are shown in Fig 11b.
Figure 11b
Fig 11b. Schematic showing singlet analysis where doublets have been gated out. Singlet vs doublets are displayed by plotting FSC-Area vs FSC-Height or FSC-Width vs FSC-Area. Alternatively, SSC signals can be used with its respective Area, height, or width signals. In this example FSC-Width vs FSC-Area produces slightly better separation of singlets. However, the scatter parameters producing the best discrimination of singlets and doublets will depend on the sample and instrument. The characteristics of the cell types in each sample and flow cytometry instrumentation such as dimensions of the laser beam may affect the optimum choice of scatter signals to use. If the discrimination of singlets and doublets is poor, test an alternative combination of parameters e.g.  FSC or SSC, Area vs height or width vs area.

4.3.2 Analysis of viability using viability dyes: Dead cells can often be difficult to distinguish using scatter alone and artifactual staining of dead cells can give inaccurate fluorescence staining of specific markers. To avoid this happening, a viability dye can be used to accurately identify dead cells and eliminate dead cells from the analysis. If possible, always include a viability dye in your analysis. Viability can be measured by flow cytometry using scatter in combination with staining of dead cells using viability dyes. Viability dyes can also be used in combination with other cell staining techniques such as immunofluorescence of surface or intracellular antigens.

There are 2 main categories of viability dyes:

(i) for staining dead cells in unfixed cells suspensions and (ii) for staining dead cells in cell suspension requiring protocols with cell fixation steps.

(i) Viability dyes for unfixed cell suspensions: These include membrane integrity dyes that are impermeant to live cells where the plasma membrane is intact but permeant to dead cells where the membrane integrity is compromised. These viability dyes can then enter the dead cells and bind to the nucleic acids in the nuclei and then exhibit increased fluorescence upon binding to double-stranded DNA (dsDNA) or RNA. Commonly used membrane integrity dyes include propidium iodide (PI), 7-aminoactinomycin D (7-AAD), DAPI and SYTOX™ Blue, Green, Orange, Red Nucleic Acid dead cell stains. The viability dye can be used in combination with other staining techniques such as surface cell analysis. The choice of viability dye must allow sufficient spectral separation of other fluorophores used in the experiment.

a. Add the viability dye just prior to flow cytometry analysis to prevent non-specific uptake by viable cells. The minimum incubation time varies with each viability dye. Staining is rapid, only taking seconds for propidium iodide and DAPI and minutes for other dyes, for example 15 minutes for 7-AAD and the Sytox dyes. Ideally, the maximum staining time is less than 30 minutes but occasionally there are unforeseen delays in running samples and the sample may be inadvertently incubated with viability dye for longer than expected. The maximum acceptable incubation time with viability dye can vary between different applications, dyes and cell types. A general rule of thumb is up to 2-3 hours for immunofluorescence analysis of various cell types such as human mononuclear cells. Therefore, if this delay is unavoidable, the data analysis should be interpreted with this issue considered e.g. a higher background staining of live cells may occur with the viability dye.

b. Avoid wash steps following staining with the viability dye because the viability dye will be washed away. However, if the cells are washed after staining with the viability dye, it must be re added prior to the flow cytometry analysis.

Fig 12
Fig 12. Histograms from ThermoFisher.
Membrane permeability nucleic acid viability dye, example showing propidium iodide
(A)   In unfixed samples, dead cells are highly fluorescent and live cells are negative
(B)   Unsuitable for fixed samples, live cells are permeabilised by fixation and separation from dead cell staining is lost.

(ii) Fixable viability dyes: These viability dyes can be used to label and discriminate dead cells for applications using protocols that require fixation, such as intracellular staining. These viability dyes covalently bind to the primary amine groups of proteins when added to a mixture of live and dead cells. The viability dyes bind to surface protein amines of both. Whereas the viability dye is permeant to dead cells as the membrane integrity is compromised and binds intracellular amines as well as surface amines. Whereas the viability dye is excluded from entering live cells. Consequently, there is higher fluorescence of dead cells compared to live cells.

Screenshot 242

Fig 13. Live and dead cell labelling with fixable viability dyes.
(A) Staining schematic is from Bio-Rad, staining either before or after fixation. This shows viable cells on the left with surface staining only for the viability dye and dead cells on the right with surface and intracellular staining of the viability dye.
(B) histogram showing fixable viability dye staining from ThermoFisher: This shows a sample containing mainly viable cells with a small proportion of dead cells. Unlike Propidium iodide and other membrane permeability viability dyes, the live cells are not completely negative with the fixable dye but shows differential staining with the more highly fluorescent dead cells.
(C) Schematic shows how the dead cells are eliminated from the analysis using a gate set on viable cells in a plot of FSC vs fluorescence of the viability dye.

There is an extensive range of fixable dead cell dyes from various manufacturers with a range of excitation and emission spectra, such as the Biolegend Zombie dyes, BD Biosciences fixable viability stains and Thermofisher fixable viability dyes. Links are provided in the list of reagents below.

4.3.3 Checking the quality of the cell suspension during data acquisition and troubleshooting problems with poor data. When all precautions and steps have been taken to prepare a single cell suspension and partial blockages still occur, samples can either be rerun or measures can sometimes be taken to eliminate poor data where the blockages have occurred.   

Using time as a parameter

fig 14
Fig 14. The diagram shows how time as a parameter can be used to check the quality of the analysis. For this kind of analysis the scatter or fluorescence of stained cells should be consistent over time. In this example gaps are produced when the sample stops running intermittently for brief periods of time. When the data acquisition is resumed the signal remains unchanged. However, a partial blockage occurs later which decreases the signal. The gate in red shows how the data can be analysed to exclude the bad data in this particular example.

Summary comments

By using a combination of techniques and reagents, best practices can be applied to achieve the optimum single cell suspension for each sample type. This will in turn improve future staining experiments and subsequently the quality of the data produced. The analysis of the cell suspension will allow you to understand how good the single cell suspension is. In addition, it will also allow data quality to be improved by gating on live, healthy, single cells and eliminating dead cells. If all else fails, data can sometimes be salvaged following data acquisition by collecting time as a parameter and gating out bad data as described.

Reagents and useful links:

Comparison of cell detachment of stromal cells, including dissociation Buffer [CDB], EDTA (ethylenediamine tetra-acetic acid) and scraping (ISCT reference):

Cell Dissociation Solution, Sigma-Aldrich #C5914, supplier Merck

Accutase, Sigma-Aldrich #A6964 supplier Merck

TrypLE, Gibco # 12604013

Cell strainers, various examples

Milenyi, different mesh sizes.

Falcon™ Round-Bottom Polystyrene Test Tubes with Cell Strainer Snap Cap (5mL), #352235

Fixable dead cell dyes, examples

Biolegend Zombie dyes.

Dead cell dyes for unfixed samples include Propidium iodide, 7AAD, DAPI, TOPRO3 and Sytox dyes

Sytox dyes (various).


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