Fees effective: June 1, 2015

These rates are for Arizona Universities. For external pricing, please contact the Facility Manager or the ARL Business Office (520-621-4064).

Acquisition has priority over data reduction.

Cell-sorting users will be charged $123.00 for setup, and then $123.00 per hour for sorting prorated to 15 minute increments.

Consultation services and the use of support equipment are available free of charge.

External customers: Please contact the Core Facility Manager.

Individual consultation services are available to discuss applications and methodologies specific to your project.

Contact the Facility Manager to set up an appointment.

There is also a small library of books and reference materials which are available for use.

Subpopulations of cells or particles that have been resolved by fluorescence or physical parameters can be isolated for further experimentation by using the instrument's cell sorting capability.

A piezoelectric quartz crystal connected to the nozzle causes the cell stream to oscillate and form droplets. The droplets are formed at a high enough frequency so that any given droplet will contain only one cell. When a cell meets the criteria defined by sort "windows", the sorting electronics system places a charge on the cell stream at exactly the time when the droplet containing the cell of interest is breaking off from the stream. The droplet passes between two metal plates that have a very high voltage.

The resulting magnetic field forces the charged droplet to be deflected toward the plate with the opposite charge. By placing a collection vessel in the deflected droplet's path, cell populations up to 99% purity can be collected at rates of approximately one thousand per second. The current flow cytometer can sort four populations at the same time.

The BD FACSAria cell sorter is operated by Cytometry Core Facility personnel only. Please contact the Core Facility Manager to discuss your cell sorting needs.

This form must be filled out and submitted to the Facility Manager prior to starting any new sorting projects involving live human or pathogenic samples.

Flow cytometry is a powerful and effective tool for analyzing the structural and functional characteristics of cells or particles in suspension. A test tube containing a single cell suspension is placed on the instrument where the sample is pushed by air under pressure through tubing where is is injected into a nozzle. The nozzle, containing sheath fluid under pressure, hydronamically focuses the cells so that they exit the nozzle in single file down the center of the stream resulting in laminar flow. The flow rate can be controlled by adjusting the differential pressure between the sheath fluid and the sample injection pressure. Laser light is then focused onto the stream where each cell is excited as it passes through the laser beam.

Photo multiplier tubes (PMT's) are detectors which collect the photon emissions from each "event" and convert them to analog voltages. The analog signals are then digitized by analog to digital converters (ADC's) and recorded as data files. Optical filters are placed before the detectors so that only wavelengths of light corresponding to specific fluorochrome emissions are collected by each detector (e.g. FITC emits in the green region therefore a 30nm bypass filter centered at 520nm could be used to collect light from this fluorochrome). Light scattered at the same wavelength and direction as the laser light, primarily from the surface of the cell, correlates with relative cell size (Forward Angle Light Scatter) while light scattered 90 degrees to the laser (Side Scatter) usually from internal structures, correlates with granularity. By correlating these two parameters, one can discriminate subpopulations of cells in peripheral blood samples, for example. Signals corresponding to cell debris or cell aggregates can also be detected and excluded from analysis on the basis of forward and side scatter.

Staining cells with multiple fluorochromes conjugated to antibodies or fluorochromes directed at other specific targets such as DNA, cytokines, or other proteins distinguishes cell subpopulations which can be quantified. Data is displayed and analyzed using histograms or two-dimensional dot plots on a computer system.

The Cytometry Core Facility has two Analyzers available for use: a two-laser BD FACSCanto II and a five-laser BD LSR II

All services being with a consultation with the Facility Staff.

Appointments for the FACSAria and the LSR II must be made by directly contacting the Facility Manager.

For access to the FACSCanto II after your initial consultation, complete and submit the New User Registration form.

1. Use your email login (the first part of your email address before the "@") as the Login Name.
2. Click Submit when you have provided all the information required. Access will not be approved until all information is complete.
3. Contact the Facility Manager when you've submitted the access request and we will respond by email with your approval.
4. Once access has been approved, you can schedule appointments for the FACSCanto II using the scheduling calendar.
5. Samples for the FACSCanto II optimally should contain 1x10⁶ cells/ml (300,000 cells minimum). For tubes that meet these requirements, allow on average 3 minutes per tube (5 minutes for cell cycle analysis) for most samples.

Experiments that look for rare events will require more time per sample.

Remember, the fewer the cells, the longer the time required for data acquisition per tube.

1. Grow cells under the desired conditions to 0.5-1x10⁷ cells/mL. Collect up to 1-2x10⁷ cells per sample. All the following steps can be performed in a single 1.7mL microfuge tube per sample.
2. Pellet the cells (a few seconds in a microfuge). Resuspend the cells in 300μl H₂O. Be sure the pellets have been dispersed before the next step.
3. Add 0.7mL 100% ethanol and vortex; incubate at 4°C for at least 30 minutes.
4. Pellet the cells as before and resuspend in 1mL of 50mM sodium citrate, pH 7.4.
5. Sonicate the cells (10 pulses of 0.9 seconds each, setting of –32 to –40).
6. Pellet the cells again and resuspend them in 1mL Na-citrate containing 0.25mg/mL RNase A. Incubate at 50-55°C for 1 hour.
7. Add 25μL of 20mg/mL Proteinase K and continue the incubation for an additional hour.
8. Pellet the cells again and resuspend the pellets in 500μl of 50mM Na-citrate. [But go instead to “Alternative” in Step 10 if your cell concentration is not very high.]
9. Prepare a batch of 1x Sytox Green (= 1μM) in 50mM Na-citrate. The stock solution provided by the vendor is 5mM = 5000x, in DMSO. Note: the manufacturer recommends wearing two pairs of gloves when handling the stock solution. The Sytox stock solution takes a while to thaw, so plan ahead.
10. Pipette out 1 ml aliquots of the 1x Sytox solution into FACS analysis tubes (Falcon 2054). Add 50μL of the cell sample (or enough to get ~1-2x10⁶cells; the goal is to have 100-200 events per second within the gated region when the samples are being read). Incubate 1 hour to overnight at 4°C; stained samples can be stored at 4°C for several weeks. [Alternative: If your sample is low-concentration, you can simply add 1 ml of 2x Sytox to the cell sample immediately after step 7, and you’ll be good to go.]
11. Use FL1 on the FACSCAN instrument. Sytox Green fluoresces more strongly than propidium iodide, so the voltage/gain won’t have to be set as high as with propidium iodide.

Sytox Green: Invitrogen, catalog # S7020

RNase A Stock Solution (10mg/mL)

• 0.2g Ribonuclease A (Sigma #R-5503)
• 200µL 1M Tris-HCl, pH 7.5
• 60µL 5M NaCl
• Bring volume up to 20mL with H₂O
• Boil 15 minutes.
• Let cool slowly to room temperature.
• Aliquot and store at -20°C.

Proteinase K Stock Solution (20mg/mL)

• Stock solution is 20mg/mL in
• 10mM Tris pH 8.0
• 1mM CaCl₂
• 50% glycerol
• To make 5mL of stock solution, dissolve 100mg Proteinase K in:
• 3.125mL 80% glycerol*
• 50μl 1 M Tris pH 8.0
• 50μl 100mM CaCl₂
• 1.775mL H₂O
• = 5mL total; dispense in 1mL portions and store at –20°C.
• *autoclaved stock; easier to deal with than 100% glycerol that’s not been sterilized

Reagents:

 This protocol has been found useful for labeling both primary cells and cell lines with the fluorescent probe CFDA-SA (carboxyfluorescein diacetate succinimidyl ester). This probe is often referred to incorrectly in the literature as "CFSE" - it should not be confused with carboxyfluorescein succinimidyl ester (the real CFSE), which is not the diacetate form and is not cell-permeable. The correct reagent can be obtained from Molecular Probes, catalog number C-1157.

Stock and Storage:

 We prepare CFDA-SE at a stock concentration 1000-fold higher than the final usage concentration (for example, 2mM if the final concentration is 2µM) in dry DMSO. Aliquot into single-usage vials and store over desiccant at -20ºC. CFDA-SE will hydrolyze quickly at room temperature in the presence of water, and much more slowly at -20ºC under desiccating conditions. Aliquoted stocks should be used for no more than 2 months. If your cells show decreased labeling with the same stock of CFDA-SE, hydrolysis is likely the cause.

Labeling Concentration and Conditions:

 Cells are usually labeled at a final CFDA-SE concentration of 0.5-5µM. The literature reports concentrations ranging from 0.2-10µM and even higher. For best results, do a titration and find the lowest concentration of CFDA-SE that will give effective cell labeling - this will vary from cell type to cell type, and also with the application. CFDA-SE labeling is somewhat toxic and can induce growth arrest and apoptosis in some cell types - therefore, it is important to find the lowest acceptable labeling concentration and check the viability after labeling. As a rough guide, 0.5-2µM is usually enough for in vitro experiments - cell tracking and generational analysis in transplanted cells may require 2-5µM. Incubation time is usually from 5-10 minutes - again, titrate to find the minimal effective conditions. We usually label in PBS or HBSS containing 0.1% BSA. All post-labeling washes should be carried out in complete media (such as RPMI with 10% FBS) - your intended tissue culture media is ideal. The high protein concentration inactivates unreacted CFDA-SE.

1. Suspend your cells in PBS or HBSS containing 0.1% BSA. Cell concentration can range widely from 1x10⁶ cell/mL (for in vitro experiments) up to 5x10⁷ cells/mL (for adoptive transfer). The cells should be in single cell suspension - if necessary, filter them through nylon mesh immediately prior to labeling. Total reaction volumes should not exceed 4mL in a 15mL tube, so prepare cell suspensions at no greater than 2mL each.
2. Prepare a solution of CFDA-SE from your DMSO stock in PBS/0.1% BSA at 2x the final labeling concentration. For example, if you are labeling at 5µM, prepare a 10µM solution. Prepare a volume of CFDA-SE equal to your cell volume above (no more than 2mL per labeling reaction)
3. Add an equal volume of CFDA-SE solution to your cell suspension. Mix gently and incubate for 5-10 minutes at 37ºC
4. Immediately fill the labeling tube to the top with the tissue culture media intended for culture (such as RPMI/10% FBS) and centrifuge. Wash the cells three times with tissue culture media at room temperature. To reduce the amount of unbound CFDA-SE in cells, we usually incubate the cells at 37ºC for 5 minutes after the second wash and prior to the third. This allows free unreacted CFDA-SE to diffuse out of the cells and be removed in the final wash.

This protocol was prepared by the Telford Lab for the NCI Medicine Branch and its friends. 11-18-00

Solutions:

Staining Buffer:

• PBS, pH 7.4-7.6
• 2% heat inactivated BCS
• 0.2% sodium azide

Fixation Buffer (4% paraformaldehyde)

• 14mL 10X PBS
• 10.8mL 37.5% formaldehyde
• 75.2mL dH₂O

Perm Buffer

• Staining Buffer + 0.5% saponin

Superperm Buffer

• 3 parts Perm Buffer + 1 part BCS (filtered)

Staining Protocol

After harvesting and any stimulation procedures, incubate cells on ice for 20 minutes. Keep cells on ice until fixation.

1. Use a minimum of 1x10⁶ cells per tube
2. If necessary, stain for surface markers as per usual FACS protocol
3. Wash cells with PBS
4. Suspend cells in 0.5mL PBS and 0.5mL fixation buffer. Gently vortex and incubate at room temperature for 20 minutes
5. Optional - To store cells after fixation, spin cells down and repeat fixation step, suspend cells in Staining Buffer and store refrigerated for up to 3 days.
6. Wash with PBS

All Intracellular Staining steps should be in saponin containing buffers

1. Wash with Perm Buffer once and then wash once with Superperm Buffer. After fixation, cells can be vortexed in the usual manner
2. Add primary antibody for intracellular staining. It is recommended to use 2x the normal amount that would be used for surface staining, but you may need to optimize by tritration
3. Incubate in the dark at room temperature for 30 minutes
4. Wash with Perm Buffer twice
5. If necessary, add secondary antibody. It is recommended to use 2x the normal amount that would be used for surface staining, but you may need to optimize by tritration. Secondary PE-conjugated reagents are recommended to get the maximum signal.
6. Incubate at room temperature for 30 minutes
7. Wash with Perm Buffer twice
8. Wash with PBS once
9. Wash with Staining Buffer once
10. Resuspend in 300-500mL Staining Buffer and analyze by FACS

Notes:

• commerical fixative and permiabilization solutions are available if preferred
• Depending on cytokine localization, stronger permeabilizing detergents such as Triton-X or NP-40 may be appropriate (0.1% - 1%)

Adapted From:

1. Shimizu Lab/Author Nadine Ottoson http://www.tc.umn.edu/~shimi002/sopicfacs.pdf
2. Think Peptides - Intracellular Cytokine Staining http://www.thinkpeptides.com/PR02TP.pdf
3. Abcam - Intracellular Staining http://www.abcam.com/ps/pdf/protocols/flow_intracellular_staining.pdf

1. Collect cells from culture or tissue. Make a single cell suspension of ~1x10⁶ cells. Wash cells by adding 3mL PBS, vortex gently and centrifuge at 200g for 5 minutes at 4ºC. Resuspend cells in Falcon tube in 50µL PBS
2. Dilute the monoclonals according to results from a previous titration
3. Add 5µL diluted monoclonal antibody to each tube. Shake tubes gently by hand. Place in an ice bath for 15 minutes in the dark
4. Add 3mL PBS to each tube. Spin at 200g for 5 minutes at 4ºC
5. Decant supernatant. Add 400µL 2% paraformaldehyde (pH 7.2-7.4) while vortexing
6. Cap tubes and store at 4ºC wrapped in foil until ready to run
    

References

Jackson, A.L., Warner Preparation, Staining and Analysis by Flow Cytometry of Peripheral Blood Leukocytes. In Manual of Clinical Laboratory Immunology. N.R. Rose, H. Friedman and H.L. Fahey, editors. American Society for Microbiology, Washington D.C., 1986, pp 226-235.