Institute of Immunology & Infection Research

Flow Cytometry Core Facility

Single Cell Preparation

Cell Preparation Guidelines for FACS analysis and cell sorting

Preparation of cells for flow cytometric analysis and cell sorting will vary depending on the source and specific requirements for each cells type. Two absolute requirements are that the cell suspension be single cell and that cells remain integrity at the end of preparation for all FACS analysis and cell sorting.

For a high-speed FACSAria cell sorter, additional requirements, including concentrating or enriching samples and preparing cells under sterile condition, may be required. The FACSAria is a high through-put, multi-laser, multi-parameter, cell sorter. Up to 11 fluorescence parameters (5 from 488 nm line, 3 from UV 407 line and 3 from 647 nm line) can be used to sort up to 4 populations simultaneously at rates of up to 40,000 cells/second.

Sample Preparation for cell sorting:

Samples should be concentrated optimally, depending on cell type.

For adherent cell lines and larger cells, which need the 100um tip on the sorter, sample should be concentrated close to 15 millions/mL.

For lymphocyte or other smaller cell types, which need the 35-70µm tip, sample should be concentrated close to 50 millions/mL.

70 micron cell strainer mesh (Falcon 352350)

40 micron cell strainer (Falcon 352340).

Cells should be resuspended in a low protein buffer (<2% FBS or BSA) in a Ca/Mg-free buffer (PBS), with or without DNase (20-100 µg/ml) to decrease the chance of clumping, typically electrotransporated B cells. Addition DNase to single cell suspension depends on your experimental endpoints. For instance, DNA cell cycle and apoptosis analysis need enough EDTA (2mM) in your sample buffer to inactivate any DNase. The addition of Hepes also may aid in viability by buffering the pH of the sample while sorting. Cell suspensions should be passed through a 40µm nylon mesh filter (available in flow lab) right before sorting. Samples and sort receptacles should be kept on ice (if applicable).

Sort Setup: As with all flow cytometry experiments, controls are absolutely critical. An unstained sample and each single stain controls, as well as FMO (fluorescence minus one, a control for which cells are stained with all reagents except the one of interest) controls, typically for multiple color analysis, are necessary for setting up the instrument. No controls, No sorting!!

If dim populations are needed for sorting, it may be beneficial to prepare FMO controls that allow you to visualize the true background including effects introduced by compensation.

Sorting Process: During the sort process, the sorter will display all sorting related parameters, including the number of cells sorted, the locations of sorted cells, abort rates and sort rates. A purity check is always performed after sorting by re-running a small fraction of the sorted population, followed the samples lines are rinsed with Bleach and Ethanol to prevent carryover from previous sorting sample.

Negative controls:

1. unstained cells (for each cell type examined)--This is absolutely required for each analysis (except DNA cell cycle). 

2. non-specific staining control for each fluorochrome or dye and each cell type (“stain” cells with either the fluorochrome isotype control, or a secondary antibody without the primary antibody)--This indicates any non-specific binding of your antibody, and is important for your interpretation of the data.

Positive controls:

-cells stained with only one antibody-fluorochrome (or other dye) at a time (for each cell type and each fluorochrome or dye)--These are essential to accurately collect and analyze data from samples stained with more than one fluorochrome or dye.

Controls are necessary to set proper compensation. 

Sample Preparation

The single most important issue for a successful sort would have to be proper sample preparation. This can be broken down into four separate components:

  1. Single Cell Suspensions
  2. Optimized Sample Concentration
  3. Proper Sort Buffer Recipe
  4. Expedient Sample Processing

Single Cell Suspensions

In order for the sorter to function properly and to be able to deliver the proper results, the cells must be in a single cell suspension, and remain that way for the duration of the sort. This becomes a more  important factor when working with adherent cell lines or tissues. Achieving a single cell suspension is the goal for a perfect sample preparation. One of the easiest tricks is to remove any large aggregates is by filtration (typically through fine 30-50um nylon mesh). There are two options. The first option, and the easiest, is to use a 12x75 mm Falcon test tube with a cell strainer cap (Falcon 352235). Simply pipet the cell suspension through the top of the cap. The second option is a little more involved. To minimize sample loss, use the techniques described below. The following tools are required (sterilize as necessary):

The next step is to aspirate the sample up through the syringe needle such that your sample is loaded along with at least 1.0mL of air as a void volume.

Once the sample has been aspirated along with the 1.0mL of void volume, carefully dispose of the needle. Maintain the syringe in an upright position so that the void bubble remains at the tip of the syringe.

The next step is to sandwich the piece of nylon mesh over the tip of the syringe using the original syringe tip. The idea is to use the tip to hold the filter mesh in place. Once the tip is in place, carefully cut off the first few mm of the tip. This will allow a hole for the filtered sample to pass through. Additionally, take care to maintain the void bubble towards the tip so no sample is accidentally lost during this step.

At this point you need to invert the needle so the void bubble rises to the plunger. Now the sample is ready to be passed through the filter mesh and the void bubble should ensure no sample remains in the syringe.

Optimized Sample Concentration

Cells must be at the proper concentration in order for the sorter to function optimally. Simply put, cells that are too concentrated will have a lower recovery due to coincidence aborts (two cell that are too close together will be rejected by the machine in order to ensure purity) and cells that are too dilute will have a longer processing time (or if they are processed faster, an increased signal CV).

Ideal Cell Suspension Concentrations

Having the sample too concentrated, or too diluted can be problematic. There is no ideal concentration that works for all cell types and sort set-ups. It is a matter of understanding some of the issues and deciding what factors are most relevant to a given cell type and experimental design.

The following is a list of concentration ranges based on machine set-up which typically correlates with cell type:

Nozzle Size Cell Types Concentration (per mL)
70um lymphocytes, thymocytes 8-15 x 106
80um activated subsets, smaller cell lines 7-10 x 106
100um larger adherent cells 5-9 x 106

These guidelines are in place to suggest where to start with, but it might be easier to tend towards a little too concentrated (towards the high end of the recommended range), but bring some additional sample buffer with you so things can be diluted as necessary.

Proper Sort Buffer Recipe

This is probably one of the most important factors to achieve an ideal sort. A properly designed buffer recipe will help maintain a single cell suspension as well as keep the cells in a good physiological state. Culture media is typically a poor sort buffer (although it can be modified).

The proper design of sort buffer for both your pre-sort sample and your collected sample is crucial for a successful sort. The following will be a basic recipe and some suggestions for modifications that might be relevant to your particular experiment. Culture media is not an ideal sort buffer for two reasons: the pH regulation fails under normal atmosphere causing the media to become basic and the calcium chloride in most culture medias is not compatible with the phosphate component of the instrument sheath buffer (the Basic Sorting Buffer without additional protein) leading to precipitation of calcium phosphate crystals. Following the suggested recipes below will help maximize the recovery and viability of your sorted cells.

Basic Sorting Buffer:

Optioal Collection Media:

For Clean Lymphoid Cells

The buffer can be simplified to HBSS with 1% FBS. The additional cations in the recipe promote better viability. Since these cells are not prone to clump, the lack of EDTA is not a problem.

For Sticky Cells

Raise the concentration of the EDTA to 5mM and use FBS that has been dialyzed against Ca/Mg++ free PBS. Some activated cells become clumpy and the chelators (EDTA) help reduce cation-dependent cell to cell adhesion.

For Adherent Cells

In order achieve good single cell preparations, one must start at the moment of detaching your cells from the plate. Typically, the trypsin (or other detachment buffer) is quenched with culture media or a PBS/FBS buffer. This is problematic because it reintroduces the cations that facilitate the cells reattaching to the plate (or each other). One must use a cation-free FBS buffer in order to stop the detachment. Additionally, the level of EDTA can be increased if necessary (but too much EDTA can be deleterious).

For Samples with High Percentage of Dead Cells

If there are a large number of dead cells in the prep, it is likely that there is soluble DNA from the dead 5 cells that will come out of solution. This DNA will start to coat the cells and and lead to severe clumping. The addition of 10U/mL DNAase II to the buffer recipe will help reduce DNA associated clumpiness.

Extremely Important: After cells are sorted into collection tubes, centrifuge cells to remove diluted buffer and replenish with fresh culture media.

These suggestions should help to optimize sample preparation for both enhanced viability and enhanced recovery. It may require some more comprehensive modifications evolving from these simplistic guidelines.

Expedient Sample Processing

The sample must be prepared in as short a time as feasible to minimize stress on the cells as sorting is a relatively harsh process. Much of this can be achieved by simplifying the  staining process and staggering the sample preps if more than one sample is being sorted.

Cell Physiology

Resting cells are typically very easy to sort, but most researchers have manipulated the system such that the cells are no longer in the most ideal state for processing. This can be addressed by setting up the instrumentation to run at lower pressures to minimize the stress on the cells. It is important for the researcher to convey any of these potential physiological issues to ensure the sort is properly configured.

End Point Requirements

The desired use for the sorted material can have a role in how the instrument is configured and how the sample can most efficiently be processed. Whether cells need to be viable, sterile or are used for DNA/RNA isolation can also have a role in instrument set-up.

Specimen Preparation

Samples must be filtered through (40 u mesh) before sorting. For example, you can use the following products: B-CMN-40 (by sq. feet) from Small Parts, Inc or Falcon 2235 tubes with the mesh in the caps from Becton Dickinson. The optimal concentration of material in the sample tubes (15-20) x10(6) cells /ml. Cells bigger than lymphocytes might need to be at lower concentrations.

The following measures are important to prevent sorting material from clumping:

  1. Concentration of fetal calf serum or BSA should not exceed 0.5%
  2. Use filtered buffer that is Ca2+ and Mg2+ free unless measurement depends on these ions
  3. Add some 0.02% EDTA
  4. Use anti-clumping agents, such as Accumax.(Innovative Cell Technologies, Inc., A Phoenix Flow Systems, Inc. Company)

It is highly recommended that unstained and single color stained controls be provided for optimum set-up.

Sufficient amount of collecting tubes (of 5ml or 15ml capacity) with media (up to 1ml or 3 ml per tube respectively) is necessary. To sort populations of low percentage, higher volume of media (up to 3 ml in 5ml tubes) is recommended for better cell recovery.

Media in collection tubes should contain at least antibiotics and 10% serum to increase viability of the cells. If many cells are expected to be collected, the serum concentration can be adjusted to100% for dilution with the sheath fluid. Wash all cells after sort and replenish with fresh media prior to placing back into culture or other experimental post sort design.


Cell concentration

In order to be able to take advantage of its high speed capabilities, cells for the MoFlo can be much more concentrated than for most other cell sorters. This is especially important for experiments where large numbers of cells are to be run.

 

Cell concentration for large cell samples: 2 to 2.5 x 107 cells/ml

This allows the cells to be sorted without having to apply excessive extra pressure to the sample tube in order to achieve a rate of 20 to 25,000 events per second.  However, cells at lower concentrations can also be sorted without problem, for example when a relatively small number of cells is to be sorted. We recommend resuspending the cells in PBS containing 2% FCS, but for very sensitive cells media with FCS can be used. 

Cell straining / filtering

In order to minimize the possiblity of nozzle clogs, it is required that the cell sample be put through a 40 micron cell strainer before sorting. These are available from Falcon (cat. no. 2340).  After filtering, the cells should be kept on ice and protected from light. For really sticky cells, it may be necessary to filter them again just prior to sorting because they can clump when sitting for longer periods of time.

 

Sample tubes

It is highly recommended that tubes to be used as sample tubes be made out of polystyrene (clear), while those for sample collection be made out of polypropylene (opaque), because of their differing electrostatic properties.  Collection tubes should be 12 x 75 mm in size, while sample tubes can be either 12 x 75 mm in size or, for large samples, 15 ml conical tubes. 

We recommend collecting cells into FCS or PBS containing FCS, especially when large volumes of cells will be sorted. The reason for this is because phosphate-buffered medium (i.e. sheath fluid) can cause precipitation of salts when mixed with carbonate-buffered medium (RPMI, MEM, etc.) leading to eventual problems in viability for sensitive cells.  To help you plan accordingly, a very general rule of thumb is that 1 x 106 sorted cells will end up having a volume of  about 1 ml after sorting. For cells which are to grow under serum-free conditions, serum-free media can be used with the collection tubes simply being changed more often. For rare cells the volume of the sorted cells won't be large enough to cause precipitation, and therefore media with serum would also be ok to use.

SORT SAMPLE PREPARATION GUIDELINES:

Do not use other type or substitute tubes!

COLLECTION TUBE PREPARATION:

SINGLE CELL SORTING:

 

Sample Preparation

Sample guidelines for the cell sort

1. Human samples must be serology tested (HIV and Hepatitis B negative) before brought to the facility.

2. Samples suppose to be single cell suspension (SCS) only (i.e. no clumps).

  • To prevent clumping during the sort, filter the SCS through the mesh (we suggest to use the 5ml Polystyrene Round-Bottom Tube with Cell-Strainer Cap, Falcon #352235);
  • You could also add 0.5 mM EDTA to your buffer solution;
  • Use 1-5% serum for the buffer.

3. All the samples to be used at the FCRC should be placed ONLY in the following types of tubes:

  • 12 x 75 mm 5ml Polystyrene Round-Bottom Tube non-sterile, Falcon #352008;
  • 12 x 75 mm 5ml Polystyrene Round-Bottom Tube with Cell-Strainer Cap, Falcon #352235;
  • 12 x 75 mm 5ml Polystyrene Round-Bottom sterile Tube with the Cap, Falcon #352054.

4. Please provide the controls - unstained and single color controls. Preferable concentration of the controls is around 1 x 106cells/ml, volume 0.5-1.0 ml.

5. The sample concentration for the sort should typically be around 10-15 x 106cells/ml. If you have fewer then 5 x 106 cells put them into the minimal volume of 0.5 ml.

6. Please provide extra 15 ml of the buffer you use for the cell samples.

7. Please provide collection tubes for the purified (post-sort) cells.

  • For the post-sort tubes use eppendorf tubes or any type of 12 x 75 mm tubes (made from glass, polypropylene, polystyrene, etc.);
  • In order to prevent cells sticking to the sides of the tubes, pre-coat the tubes, filling them with the serum for 30 minutes before the sort;
  • For the sterile sort the post-sort tubes should be also sterile.
Murine Tissue
Optical Platform

Preparation of single cell suspensions from mouse tissues

 

For collection of blood the heart must still be beating therefore mice have to be anaesthetised. If peritoneal exudate and thymus are to be collected it is best to kill mice using anaesthesia or CO2, as cervical dislocation can cause bleeding which contaminates these tissues, especially with red blood cells, making flow cytometry analysis difficult.

 

Peripheral Blood

 

As the heart has to be beating for blood sampling it should be collected in accordance with home office project and personal licence procedures.

 

If there is no requirement for recovery post-anaesthesia Euthatal (available in BRF) can be used.

 

For collection of both plasma and cells for analysis, whole blood should be collected into an eppendorf tube containing 10IU/ml heparin

 

Isolation of plasma

  • Spin sample in microfuge at 5 000rpm for 4minutes
  • Carefully transfer plasma to fresh eppendorf using micropipette fitted with yellow tip – store at minus 80 C until required
  • Retain pellet for isolation of isolation of white blood cells (leukocytes)

Plasma can be used to assess cytokine/chemokine levels by ELISA/Cytokine Bead Assay

 

Isolation of white blood cells (leukocytes)

It is necessary to remove the majority of the red cells from the sample to allow detection of the white cells especially for flow cytometry analysis of stained leukocytes. There are a number of ways to do this, the cheapest, fastest and most reliable is hypotonic shock

·          Following isolation of plasma loosen retained pellet by flicking base of tube

·          Add 0.5ml PBS, mix by aspiration and transfer to 15ml conical tube

·          Rinse eppendorf twice more with 1ml aliquots of PBS and pool into 15ml conical

·          Add PBS to give a final volume of 12ml

·          Centrifuge at 450g for 5 minutes

·          Discard supernatant

This gives a loosely packed pellet and the supernatant therefore has to be removed and discarded by aspiration using a Pasteur pipette.

The sample is now ready for red cell removal by hypotonic shock. Before starting have the following to hand

§         Distilled water

§         10x PBS

§         1ml micropipette set at 1ml with tip attached

§         10ml pipette

§         Electronic pipettor

·          Loosen pellet by flicking base of tube

·          Add 9ml of distilled water, cap tube and mix by inversion for 12 seconds

During this process the cell suspension should go from opaque red to clear red

·          Add 1ml 10x PBS cap tube and mix by inversion

·          Centrifuge at 450g for 5 minutes

·          Discard supernatant

The volume of the pellet is considerably reduced by this point, however, red cell ghosts remain and have to be removed by washing. As above, the pellet is loosely packed, so the supernatant has to be removed and discarded by aspiration using a Pasteur pipette.

To remove red cell ghosts

·          Loosen pellet by flicking base of tube

·          Add 10ml of PBS,

·          Centrifuge at 450g for 5 minutes

·          Discard supernatant by aspiration

·          Resuspend pellet in 0.5ml PBS

Count cells – as blood is still rich in red blood cells even after hypotonic shock and the number of white cells need to be counted, it is best to dilute the aliquot for counting in white cell counting fluid (WCF 3% acetic acid in distilled water plus a @knife point of Gentian or crystal violet). Usually 10ul into 40ul WCF gives a reasonable count on a haemocytometer. The count should be done as soon as possible after dilution as though the red blood cells are destroyed immediately, eventually the white cells will be destroyed too.

·          For staining of cells for flow cytometry cells should be resuspended at 1 x 107/ml in FACS-PBS.

 

Peritoneal exudates cells (PEC)

 

1.      Lay mouse on its back

2.      Peel back skin to expose peritoneal membrane taking care not to puncture the membrane

3.      Fill 5ml syringe with PBS and attach a 21g needle (green)

4.      Insert the needle through the peritoneal membrane into the peritoneal cavity cavity low on the mouses’s left hand side

5.      Remove needle from peritoneal cavity and gently shake mouse by holding hind feet

6.      Using the same syringe and needle recover PBS from the peritoneal cavity

7.      Repeat steps 3-6 twice more

It is impossible to recover the complete volume at any wash step and usually recovery is lowest from the first wash step

 

8.      Make cells up to 25ml in PBS

9.      Centrifuge at 400g for 5 minutes (about 1600rpm in bench centrifuge)

10.  Discard supernatant

11.  Loosen pellet by flicking base of tube

12.  Resuspend in 1ml PBS

13.  Count cells – unless there has been bleeding into the peritoneal cavity there is no need to use WCF, cells can be counted without dilution

14.  For staining of cells for flow cytometry cells should be resuspended at 1 x 107/ml in FACS-PBS.

Bone Marrow

 

This is usually isolated from the long bones – femur (hind leg) and humerus (fore leg).

·          Peel back skin to expose long bones

·          Using a pair of curved scissors trim away most of the muscle from the bone

·          Holding the adjacent bone (tibia or radius) with forceps, reflex the joint (knee or elbow) and cut across the upper and lower joint to remove the required bone.

·          Place bone in sterile PBS in petri dish for storage until all required bones have been collected

Marrow is extracted from the bones by flushing using a syringe and needle. It is best to use a syringe which will hold the total volume of marrow generated during flushing for the aspiration step at the end of preparation at all stages avoid foaming

·          Fill a 2ml syringe with PBS and attach a 25g (orange needle)

·          Holding bone with forceps use scissore to remove the epiphyses (ends) from the bone

·          Insert the needle into one end of bone and flush through into universal tube with PBS

·          Repeat procedure through other end of bone

 

At this point the extracted marrow will be a mix of single cells, clumps and ‘tubes’, to produce a single cell suspension the extracted marrow should be aspirated through a series of needles of increasing gauge.

·          Draw up through a 19g needle (white)

·          Expel through a 21g needle (green)

·          Draw up through a 23g needle (blue)

·          Expel through a 21g needle (orange)

 

The single cell suspension is now prepared

·          Make cells up to 25ml in PBS

·          Centrifuge at 400g for 5 minutes (about 1600rpm in bench centrifuge)

·          Discard supernatant

·          Loosen pellet by flicking base of tube

·          Resuspend in 2ml PBS

·          Count cells – as bone marrow is rich in red blood cells and the number of white cells needs to be counted, it is best to dilute the aliquot for counting in white cell counting fluid (WCF 3% acetic acid in distilled water plus a @knife point of Gentian or crystal violet). Usually 10ul into 40ul WCF gives a reasonable count on a haemocytometer. The count should be done as soon as possible after dilution as though the red blood cells are destroyed immediately, eventually the white cells will be destroyed too.

·          Resuspend cells at 5 x 107/ml in PBS for injection. For staining of cells for flow cytometry cells should be resuspended at 1 x 107/ml in FACS-PBS.

 

 

Spleen

 

·          Having peeled back the skin for removal of bone marrow and harvested PEC

·          Lay mouse on its back

·          Using forceps pick up peritoneal membrane near the midline just below the sternum and make an incision. Then cut along below the diaphragm and  then down towards the tail to allow the peritoneal membrane to be pulled back to reveal the abdominal contents. The spleen is on the mouse’s left side tucked under the liver.

·          Grasp the end of the spleen and lift it upwards, then release the spleen from the  fat and connective tissue using scissors. Keep spleen in PBS until ready to prepare a single cell suspension.

·          Before processing remember to weigh the spleen.

 

There are a number of ways to prepare a single cell suspension of spleen the quickest and simplest uses a 5ml loose fitting glass homogeniser.

·          Use forceps to transfer spleen into homogeniser

·          Add 1ml PBS

·          Insert glass rod into homogeniser, exert gentle downward force and turn rod 3 times – this should be sufficient to release the pulp from the spleen capsule

·          Add 4ml PBS to homogeniser and mix by gentle aspiration, avoiding foaming

·          Debris and clumps can be removed by allowing the cell suspension to stand for 2-3minutes and decanting the suspended cells to a fresh tube by pipetting or by passing the cell suspension through nylon mesh.

·          Make cells up to 25ml in PBS

·          Centrifuge at 400g for 5 minutes (about 1600rpm in bench centrifuge)

·          Discard supernatant

·          Loosen pellet by flicking base of tube

·          Resuspend in 5ml PBS

Count cells – as spleen is rich in red blood cells and the number of white cells need to be counted, it is best to dilute the aliquot for counting in white cell counting fluid (WCF 3% acetic acid in distilled water plus a @knife point of Gentian or crystal violet). Usually 10ul into 40ul WCF gives a reasonable count on a haemocytometer. The count should be done as soon as possible after dilution as though the red blood cells are destroyed immediately, eventually the white cells will be destroyed too.

·          For staining of cells for flow cytometry cells should be resuspended at 1 x 107/ml in FACS-PBS.

 

 

Thymus

·          Having peeled back the skin

·          Use forceps to grasp xiphoid sternum and using scissors cut along and above the diaphragm to gain entry into the thorax

·          Use scissors to cut up either side of the rib cage

·          Grasp xiphoid sternum with forceps and lift up and reflex over head

·          Thymus overlies the heart

·          Use forceps to release thymus from heart

·          Then use a pair of curved forceps to grasp the thymus at its origin and use a second pair of forceps placed just below the first pair to detatch the tissue

·          Keep in PBS until ready to prepare a single cell suspension.

·          Use forceps to transfer thymus into homogeniser

·          Add 1ml PBS

·          Insert glass rod into homogeniser, exert gentle downward force and turn rod 3 times – this should be sufficient to release cells from the capsule

·          Add 4ml PBS to homogeniser and mix by gentle aspiration, avoiding foaming

·          Debris and clumps can be removed by allowing the cell suspension to stand for 2-3minutes and decanting the suspended cells to a fresh tube by pipetting or by passing the cell suspension through nylon mesh.

·          Make cells up to 25ml in PBS

·          Centrifuge at 400g for 5 minutes (about 1600rpm in bench centrifuge)

·          Discard supernatant

·          Loosen pellet by flicking base of tube

·          Resuspend in 5ml PBS

·          Count cells –unless there has been bleeding into the thorax there is no need to use WCF, cells can be counted without dilution

·          For staining of cells for flow cytometry cells should be resuspended at 1 x 107/ml in FACS-PBS.

 

 

FACS-PBS

 

PBS supplemented with

0.1% BSA

0.1% sodium azide

 

This is highly toxic and should not be ingested