FAQs - Sample preparation
Which flow cytometer should I book?
For any questions about your experimental design and which machine will best suit your needs, it is recommended that you ask a FlowCore staff member directly.
However there are a few restrictions...
Do you need the UV laser?
In the case of sorting, the Influx 1, Influx 3, and ARIA Fusion have a UV laser - when booking you must specify that you need the laser as it is not routinely used.
For analysis; LSRIIb, Fortessa X20b and X20c have UV lasers.
Are you sorting biohazardous material?
YOU MUST CONTACT FLOWCORE IF YOU REQUIRE SORTING OF BIOHAZARDOUS MATERIAL
All primary human samples or potentially infectious samples must be booked on the Influx 1, Influx 3, or the Aria Fusion. Please contact a FlowCore staff member if you are unable to find a suitable timeslot.
What instrument setup do I need?
As a general guide;
Small cells (eg. lymphocytes from primary tissue or blood) are suitable for sorting with the high-speed 70um / 60psi setup. Maximum sorting speed for 70um / 60psi is ~ 25,000 cells/second.
Large cells (eg. hepatocytes, epithelial cells, tumor, cell lines) require the larger and more gentle 100um / 20psi setup. Maximum sorting speed for 100um / 20psi is ~ 5,000 cells/second.
How do I decide which fluorochromes to use?
Wherever possible you should use fluorescent dyes and proteins with as little spillover as possible between them. This will minimise the data spread introduced by these spillover effects and, in turn, maximise the sensitivity of your assay and accuracy of your measurements.
To achieve minimal fluorescence spillover:
Use fluorochromes which are excited by different lasers, eg. a good basic four colour combination would be:
Blue laser - FITC
Violet laser - Pacific Blue
Yellow-green laser - PE
Red laser - APC
Use fluorochrome combinations with as large a difference in emission wavelength as possible (eg. PE-Cy7 or APC-Cy7 could comfortably be added to the list of dyes above).
To assist with choosing well separated fluorochromes, BD has a spectrum viewer available on their web site (see link below). You can use these to match your fluorochromes to the lasers and filters in the flow cytometer you plan to use, as well as visualising the amount of spillover to expect.
If you only need a few colours in your assay, we recommend choosing the brightest fluorochromes available (typically the large protein dyes, i.e. PE and APC). In descending order of brightness, here is a list of recommended fluorochromes:
APC / Alexa647
FITC / Alexa488
Pacific Blue / BV421
Tandem dyes such as PE-Cy7 are highly susceptible to oxidation (which will cause it to fluoresce increasing at the PE emission wavelength rather than the Cy7 emission wavelength). It must therefore be treated extremely well, minimising the time it is out of the fridge and exposed to light.
Ensure that you are not using fluorochrome combinations with similar or identical emission wavelengths (remembering to consider the viability dye as well). Some examples of incompatibility include:
Pacific Blue and DAPI
RFP and PI
APC / Alexa647 and PE-Cy5
PE and Alexa568 / 594
Do I need a viability dye?
Whenever possible, yes!
Dead cells cause nothing but trouble in flow cytometry data - they're sticky (and will non-specifically bind your labelled antibody) and tend to have higher autofluorescence, leading to false positive signals.
Viability dyes such as propidium iodide (PI), 7AAD, and DAPI allow you to discriminate between live and dead cells and thus ensure only viable cells are analysed or sorted for further experimentation. The only time you shouldn't use a viability dye is if you have no fluorescence channels to spare for this in your assay.
There are many viability dyes available (in addition to the ones mentioned above), to complement most staining panels.
What controls should I bring?
As with any experiment it can only be as good as your controls! They must be prepared for each experiment as the set up of the machine will vary from day to day. Controls for flow cytometry are are important as they are confusing, so do not hesitate to ask a FlowCore staff member if you need help designing your assay.
The minimum controls required are:
1. Negative control
- - also known as unstained or untransfected
- -may contain a viability dye
- - used to set voltages for scatter and fluorescence channels
2. Single colour controls
- - only have one fluorescent parameter per control
- - used for calculating fluorescence spillover to other channels
- - must be as bright or brighter than staining in your test sample
- - with the exception of tandem dyes (which vary from batch to batch) you can use a different antibody for this control
- - you can use antibody-capture beads for this control
Other useful controls:
- - secondary only (where you are using two-step staining)
- - isotype control (accounts for false positives due to antibody isotype)
- - Fluorescence minus one, FMO, (for setting boundaries between positive and negative with certainty - very useful when sorting rare or dim populations.)
How can I prevent my samples from clumping?
Even the cleanest of cell sample preparations will have a degree of cell clumping, which is why all samples must be filtered prior to being run on FlowCore equipment.
Sometimes you will find that your cell sample clumps even after filtering. This is particularly common with adherent cell lines and samples from digested tissue, i.e. where enzymatic and/or mechanical dissociation is necessary. It is important to be as gentle with your cell preparation as possible. Clumping is usually caused by DNA being released into solution; by minimising the amount of mechanical digestion you will reduce cellular rupture due to shearing forces, and thereby reduce the amount of DNA in solution.
Some other tricks you might try:
- - adding EDTA to your final suspension (1 mM)
- - adding DNAse to your final cell suspension
- - diluting your cells - sometimes there is a critical concentration below which they are less likely to clump
What is the best way to filter my cells?
No matter how clean a sample preparation is, there will inevitably be a degree of cell clumping which has the potential of causing blockages in the fluidics of the flow cytometers. At best a blockage (or even a partial blockage) will cause distortions in your data, at worst it will require half an hour to clear and re-calibrate the machine, which may mean you run out of time before finishing your assay.
For this reason we require that you filter all of your samples immediately prior to being run on FlowCore equipment.
Here are just a few filtering options depending on your sample size and sterile requirement:
5 ml polystyrene round-bottomed tubes with cell strainer cap (35 um, sterile)
- - good for volumes up to ~ 3 ml
- - use to filter directly into either polystyrene or polypropylene tubes (required for cell sorting)
- - available from the Medicine Store
Nylon mesh strainers
- - 40 um, 70 um, 100 um
- - these fit onto 50 ml centrifuge tubes
- - good for filtering cells in bulk at beginning of experiment if you have a large volume
Nylon Mesh, NITEX, 50 um (31% open, 153 cm wide)
- - can be cut to the size you require
- - good for analysis samples that do not need to be sterile
- - can filter directly into plates or tubes
- - order from Sefar Filter Specialists Pty Ltd
At what concentration should I supply my samples?
As a general rule 1x107 cells per mL is a good starting point, but the best concentration at which to suspend your cells is dependent on the nature of the sample.
Samples that require little or no dissociation (e.g. peripheral blood, bone marrow, lymph node, non-adherent cell lines) can be suspended as high as 2x107 cells per mL.
Samples prepared by dissociation, especially those likely to be "sticky" (e.g. brain, liver, kidney, adherent cell lines), should be more dilute to minimise clumping and thus reduce the chance of causing blockages in the flow cytometer. Such samples could be suspended at 3-5x106 cells per mL.
If in doubt, suspend samples in a higher concentration - we can easily dilute the sample to a lower concentration. However, please keep in mind the following minimum required volumes for running samples:
When using the high-throughout plate sampler on the analysers, this will be the "dead" volume for the plate type you are using (typically 50uL, but ask a FlowCore staff member if in doubt).
For all other applications: 200uL.
What should I supply my samples in?
Unless you are planning to use the high throughput plate sampler on the LSRIIa or LSRIIb, all samples should be supplied in either the "hard" (polystyrene) or "soft" (polypropylene) 5ml FACS tubes, depending on which machine you are using:
The LSRII and Fortessa analysers require polystyrene (hard) tubes
The Influx cell sorters require polypropylene (soft) tubes
The Aria Fusion is able to run both polystyrene and polypropylene tubes.
If you are using the high throughput plate sampler (HTS) on the LSRII, please make sure you use plates manufactured by BD - some plates from other manufacturers have slightly different dimensions, using these plates on the HTS will damage the automated sampling probe.
BD manufacture all these tubes and plates - the catalogue numbers are listed in the table below.
Most of these items are available from the Medicine Store; located on the ground floor of building 77.
How do I gain entry to FlowCore?
Swipe card access to the facility is available for holders of a Monash University staff/student card and for CSIRO employees with FOBs. You will need to have completed your facility access registration, and you will also be required to complete a Local Area Safety induction before your access can be approved.
To request swipe card access please email email@example.com with the following details:
staff/student ID number
How do I make a booking?
Bookings for FlowCore equipment can be made via the online calendar
Bookings made three weeks in advance are subject to change by negotiation.
FlowCore reserves the right to change booking times or instrument when necessary, but will endeavour to give researchers as much notice as possible.
How much time should I book?
Sorting time is dependent on the concentration and volume of your cell suspension and, for the cell sorters, the nozzle size and sheath pressure.
Your best bet is to consult a FlowCore staff member and be generous with your booking time if you're running an assay for the first time. As a general guide:
For small cell types (blood cells etc.) we can run up to 25,000 cells through the instrument per second; assuming your sample is sufficiently concentrated.
For larger cell types (cell lines, large primary cells eg. hepatocytes), the maximum event rate will be closer to 5,000 cells per second.
The LSRII is capable of running cells at up to 30,000 cells per second. Again; this depends on your sample being concentrated enough.
Don't forget to take into consideration the time it takes to change tubes ... this can take much longer than you expect if you have a lot of samples!
Please also note that there may be a nozzle change between bookings, and therefore the recalibration time will be shared between the 2 bookings.
- - Ie. Booking 1 will finish 15mins early, and booking 2 will start 15mins late; allowing 30mins for nozzle change and recalibration.
How many populations can I sort?
You can sort up to 4 different populations simultaneously on the Influx 1, Influx 2 and Aria Fusion.
The Influx 3 is capable of sorting up to 6 populations simultaneously, however some cell types may not be suitable for 6 way sorting, due to side-stream instability. Please contact FlowCore if you are considering a 6-way sort.
What can I use to collect my sorted populations?
When collecting into tubes, the following formats are compatible with all of the cell sorters:
5ml FACS tubes (polypropylene / 'soft')
10-15ml centrifuge tubes
50ml centrifuge tubes
Any tissue culture plate format
You can use any buffer/media suitable for your cells in the collection tubes
Please inform us if you are sorting into media containing a cell lysing agent