Advice and guidelines

• Discuss the sort well in advance with the cell sorter operator. In order to prepare for the sort, the operator will need to know the stain you will be using; the kind of cells to be sorted; the proportion of your desired cells in the original cell suspension; the number of cells per sample you will bring to the facility; the number of samples; whether you want to sort into tubes or plates; what sheath fluid will keep your cells happy; does your sort need to be sterile or not, etc...
• Cells must be in a single cell suspension. Use cell strainers to filter your cells if you are handling clumpy samples.
• If you can, bring to the flow lab twice the number of unsorted cells that you calculate are needed to provide you with enough of your sorted population. Do everything you can to enrich your cells before you get to the flow lab, this will save you time and money.
• Bring an extra control tube with enough unstained cells if you want to sort a population based on fluorescent protein levels or on any fluorescent dye. This will allow us to draw a lign between negative and positive cells. There is no minimum number of unstained cells that you should bring with you but 50,000 to 300,000 unstained cells is ideal.
• Your cells should be made up to a concentration of about 20-40 million per ml of PBS in less than 3% serum (more proteins make the stream unstable and can clog the sorter) in a buffer that is less sensitive to pH change (10mM HEPES, pH 7.2) and that contains 10ug/ml DNAse I and 1-4 mM EDTA to help cells stay in single cell suspension and to prevent clumpy and sticky adherent or dying cells from clogging the cell sorter
• Always keep your cells on ice to stop cell death.
• Plan your time by remembering that your sample cells will be flowing through the cell sorter at a rate of about 20 to 100 million per hour, depending on sample quality, cell size, desired purity and recovery.
• Bring your collection tube(s) of choice (15 or 50 ml Falcon, FACS, or Eppendorf tubes) or a plate containing medium/serum (3 ml per Falcon tubes, 0.5 ml per FACS tubes and 0.2 ml per 96-well plate) into which the cells will be sorted. If you expect to collect very few cells (less than 100,000), it's better to use Eppendorf tubes containing 1ml of your preferred collection medium. We cannot sort into 15 or 50 ml Falcon tubes if you are collecting more than 2 cell populations.
• Sorted cells will be at a concentration of 200,000 to 1 million cells per ml depending on the micro nozzle fitted on the sort head. If using FACS tubes, bring one tube for every 3 million sorted cells (plus two extra in case of problems) when sorting small cells (size up to 15 um ) and one tube for every million sorted cells when sorting big cells (size higher than 15 um).
• If possible, re-run a few of the sorted cells at the end of the sort to check their purity. The sort operator would also appreciate feedback about how many sorted cells you count and about their viability (and sterility, if relevant).
• If you are cloning your cells, prepare 96-well plates in advance by adding 200 to 250 microliters culture medium to each well and bring plates with you to the facility. Most single cells don't like to be alone in a well by itself. To help a single cell form a colony mix conditioned culture medium* with fresh culture medium in 1:1 ratio. Cells can be cloned either based on scattered light (no staining is require) or based on fluorescence from a cell surface markers defined cell population. We can sort up to 40 plates per hour depending on sample quality, number of samples, and frequency of cell population of interest. Spinning the plates after single cell cloning helps adherent cells settle down.

 *old culture medium where same type of cells have grown for a period of time that is sterilised by filtration through a 0.22 um sterile filter.

Collecting sorted cells for RNA extraction

Cell sorting:
RNA extraction works better when cells are sorted alive (non-fixed). Extracting RNA from fixed cells is difficult but not impossible. If you are fixing samples prior to cell sorting then everything must be prepared in an RNase-free environment using RNase-free buffers and Eppendorfs or FACS-tubes. Adding RNase inhibitors to all buffers, including collection media, and cell suspensions, will help keep all media free of active RNases.
Good quality samples with limited amounts of debris, plenty of healthy cells and no clumps will improve the yield of collected cells. Any pre-enrichment steps can also help.
Use of a viability dye will also improve the quality of cells collected by eliminating dying cells; dying cells in your collection tube can interfere with the RNA extraction process. Viability dye can be added to live cells at the facility immediately before the sort - we provide non-sterile DAPI, PI and ToPro3 for this purpose. Dead cells can also be excluded during cell sortin of fixed cells.

Collection media for RNA extraction:
There are different methods of collecting cells for RNA extraction:
1) Collect cells into tubes of pure serum, media or a PBS based buffer. The collected cells are kept on ice until they can be spun down to a pellet. The pellet can either be frozen or immediately used for RNA extraction. Some cells will be lost during the spinning process but any excess of PBS added to the collection tube by the cell sorter will be removed. This method works better when cells are robust or when collected cell number is not a limiting factor.
2) Collect cells into tubes containing RNA extraction buffer. The collected cells are then normally vortexed and put on dry-ice to snap freeze for storage before extracting RNA. The cell sorter will add a volume of sheath fluid to the collection tube which will dilute the RNA extraction buffer and could alter its efficacy. The volume added will depend on the nozzle size of the cell sorter; for every 100,000 cells collected:
70-micrometre nozzle: 100uL of sheath fluid (PBS plus preservatives) is added to the collection tube (1nL per cell).
85-micrometre nozzle: 200uL of sheath fluid (PBS plus preservatives) is added to the collection tube (2nL per cell)
100-micrometre nozzle: 330uL of sheath fluid (PBS plus preservatives) is added to the collection tube (3.3nL per cell).
130-micrometre nozzle: 660uL of sheath fluid (PBS plus preservatives) is added to the collection tube (6.6nL per cell).
A limited number of cells can be assigned to a collection tube to avoid diluting the extraction buffer too much and/or to keep within the limits the extraction kit is designed for.
3) Collect a single-cell per well into 96 or 384-well plates containing RNA extraction buffer for single-cell sequencing. The plate must be spun down briefly to ensure cells are immersed in the buffer before the extraction protocol or freezing at -80C.

Troubleshooting:
RNA extraction after sorting is not always straightforward and a fellow member of your lab may have perfected the technique already. Speak to colleagues and staff about any issues preferrably before you plan your sort. A few common problems are below.

Limited number of cells collected
Debris, clumps, dying cells and red blood cells are just a few factors that will affect the efficacy of the cell sorter to successfully sort the target population. Better preparation of the sample is needed to increase the yield.

No RNA quantified in NanoDrop
Users with a small number of collected cells may not see any trace of RNA when measuring on NanoDrop. This is usually due to very low levels of RNA in the sample beyond the limit of detection. Several users reported this and were still able to amplify usable cDNA from their samples (from as little as 14 collected cells!).

No pellet of cells in collection tube
Cells collected in PBS, media or serum-based buffer need spinning to a pellet before freezing or resuspension in extraction buffer. If this is the case, has the correct g-force been used in the centrifuge? To make sure that your cells are sorted and collected properly, we usually run a few microliters of collected cells back through the sorter as a 'purity check' to confirm presence of the correct cells in the collection tube.

Theoretical limitations

Sorting is an art, a skill, and a science. It is, at all three of these levels, a working partnership between the research user and the cell sorter operator. There are certain theoretical limitations that a user needs to understand about the technique itself and there are also certain things that a user can do to maximise their chances of sorting the required numbers of desired cells in good condition for further manipulation.

Cell number

Although flow sorting may seem magical, it cannot create life: You will never end up with more cells than you started out with. If your initial sample has a million cells and you want to sort 10% of that population, then you cannot get back any more than 100,000 cells at the end of the sorting procedure (unless your counting method is not reliable). In fact, there are certain built in "abort" conditions, so that your theoretical maximum could be only 90% of this value. (Abort rates rise when sorting at high speeds, sorting relatively rare cell populations, or sorting poorly prepared samples). You must make sure that you start out with enough cells to cover your requirements and some extra to cover abort losses (experienced sorters usually try to start out with at least twice the number of cells that they need - just to be safe).

Time

The cell sorters FACSAria III and MoFlo XDP can sort at very high flow rates. they can both perform cell sorting up to an amazing 70,000 events/sec, but with sort efficiency decreasing with high flow rates. The cell sorting speed is also limited by the quality of cell preparation, which affects the abort rate, and it can be adjusted depending on the number of cells to be sorted and the time available. The cell sorter is usually set to process cells at an abort rate of less than 10%. In order to give higher flow rate, the unsorted cells should be at a concentration of about 20 to 40 million per ml. When cells flow at a rate of 30,000 per second, the total number of your original cells that will be processed each hour is about 100 million. If, however, your desired cells are 50% of the total, then you will get back about 50 million cells per hour. If your desired cells are 10% of the total, then you will get back about 10 million cells per hour. If your desired cells are 1% of the total, then you will get back about 1 million cells per hour. See Table below for the specific amount of time you will need to get an appropriate cell number back. This information may help you plan your time. It may also help you plan your budget - and may convince you to do anything you can to enrich your cell population before you bring your sample to the flow lab. Aside from financial considerations, it is also true that the less time you spend sorting, the more viable your sorted cells will be.

Sort time, given an initial positive fraction and a desired amount of cells of interest, when the cell sorter is running at 30,000 events/sec and 100% efficiency.

Hints

Recovery/Viability

After an effective sort, it is a pity to lose precious cells as a result of careless post-sort handling. Bearing in mind the theoretical limits mentioned above, cell recovery and viability after the sort event can be maximized by some simple procedures. Especially when sorting rare populations, it is important to sort cells into tubes that contain a small volume of appropriate medium in order to keep the cells happy and viable after they have travelled through the laser beam and been deflected to the left or right. For blood cells, it is probably best to sort into about 0.5 ml or more of 100% serum. It is important to realize that the cells will get diluted into cytometer sheath fluid (usually PBS) during sorting. Different types of cells have different preferences. You need to decide on the appropriate serum or medium into which you want to sort your cells. It is also possible to change the sheath fluid if your cells do not like PBS. It is up to the user to consider the cells' preferences and discuss these with the sorting operator in advance.

If you are sorting into FACS tubes, you should therefore bring with you to the flow lab these tubes containing about 0.5 ml of serum/medium and enough of those tubes to collect all the sorted cells. Bring one FACS tube for every 3 million sorted cells. If you are sorting into Falcon tubes, bring with you these tubes containing about 3 ml of serum/medium - and enough of those tubes to receive all your sorted cells. Bring one 15 ml tube for every 3 million cells you plan to collect (and few extra tubes just to cover any problems that may occur). We cannot use Falcon tubes when doing a 4-way sort. If you are sorting into wells of a 96-well plate, put into each well 100-200ml culture medium where cells will be happy for few days. Another factor to remember is that the cells to be sorted (and the cells that have already been sorted) are not being kept under carbon dioxide. Therefore use a HEPES or similar buffer, but avoid bicarbonate buffer as it will not maintain an appropriate pH for long. As mentioned above, a final factor is time: the more you can do to pre-enrich cells before sorting, the faster they will be sorted and the happier they will be.

Sterility

Although the flow sorter is a cumbersome beast, it can be sterilized to provide the user with sorted cells for long term culture. The sort operator needs to be informed in advance about a requirement for sterility since flow lines, tubes, and filters all need to be specially treated. Because there is no such thing as absolute sterility (and effective sterility will vary depending on the antibiotics present in your culture medium), if you intend to grow sorted cells for a long time without much antibiotic cover, you may want to use a short test sort at some time before a big experiment to check flow sterility under your own culture conditions.

Purity

Cell sorting will, routinely, produce populations that are 95-99% pure for the desired cells. Apparent lack of purity can result from various reasons that have nothing to do with the effectiveness of the sorting procedure. For example, purity will be less if the sorted cells are not a clearly discrete population and overlap with unwanted cells. Also, clumps of cells that separate after sorting may give the appearance of a sorted fluorescent population contaminated with non-fluorescent cells. Loss of viability after sorting may result in cells with different scatter properties from those of the original cells. Sorting cells away from platelets/debris is often not effective because platelets and debris stick to sorted cells and fall off after the sort. Capping of surface markers after the sorting procedure will often result in sorted cells that are somewhat less fluorescent than the original selected population. But, with these limitations, we can expect to take cells that are, for example, less than 1% in an original sample, and end up with about 98% purity. However, you shouldn't take this for granted. At the end of every sort, you should run an aliquot of the sorted cells through the cytometer in order to check their purity in comparison with the original, pre-sort sample.

Controls

Cell sorting is a further elaboration of the flow technique used for cell analysis. Controls are, therefore, just as important for sorting as for non-sorting flow cytometry. If you want to sort fluorescent cells, then you need to run control or unstained cells first in order to see where to "draw the line" between the fluorescent and non-fluorescent populations. What this means is that you will have to bring a tube of about 300,000 unstained cells (with experience you will need far less, usually 100,000) in order to assist the sorting operator in setting the cytometer to make its sort decisions. If you are staining your cells with more than one colour, you will need to add extra controls. You should usually provide one single colour control for each colour you are using. These control cells should, of course, be sterile if you are doing a sterile sort (so that they will not contaminate the clean cytometer). If your cells are precious and you can't waste them to prepare single colour controls, there are alternatives that use beads and that can help you calculate a compensation matrix that will be valid for your multicolour stained samples. For more information, please come and talk to us.

Cell analysis guidelines

Data Analysis Software

UCL ICH Flow Cytometry Core Facility users have several data analysis programs at their disposition:

• FACSDiva, FACSArray and FCAP software from Becton Dickinson
• Summit, CytExpert and Kaluza software from Beckman Coulter
• FlowJo from TreeStar, for universal Flow Cytometry Standard (FCS) files analysis.

Dedicated Mac and PC computers running FlowJo, Summit, ModFiT, Kaluza and FACSDiva analysis software are available for free to all facility users. The facility also offers FCAP array software for CBA analysis.

Guidelines for a valid flow cytometry experiment

With the rapid advances in flow cytometry technology, new fully digitized instruments are replacing the old analog machines and new software are developed. To understand and correctly interpret flow cytometry data, it is becoming essential to know how these data are generated and recorded. There are few papers that explain these important advances and outline guidelines as to how to record and report flow cytometry data.

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  facilities-flowcyt-miflowcyt_standard-isac_recommendations_copy.pdf
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  facilities-flowcyt-optimizing_a_multi-colour_immunophenotyping_assay.pdf
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  facilities-flowcyt-selecting_reagents_for_multicolour_immunophenotyping_assay.pdf
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  facilities-flowcyt-interpreting_flow_cytometry_data-a_guide_for_the_perplexed.pdf

Biosafety

The following are important health and safety documents for flow cytometry users:

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  facilities-flowcyt-biosafety_standard_for_sorting_of_unfixed_cells_2007.pdf
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  facilities-flowcyt-introduction_to_the_biosafety_guidelines_.pdf
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  facilities-flowcyt-biosafetysectionforhivimmuniophenotypingqa.pdf

The facility publishes its own risk assessment that is sent to each user when they book an induction with the facility manager.