Date of revision and approval

WIMR Document reference: WIMR-SWP-OP-WS-501.02

Date of approval: TBD

Approval authority: Scientific Platforms Manager, Workplace Health and Safety Manager, Advanced Specialist Flow Cytometry

Functional unit: Flow Cytometry

Enquiries contact name: Xin Wang, Suat Dervish, Edwin Lau

Enquiries contact: wrhflow@sydney.edu.au

Objective

This document describes the operational setup and operation for the BD FACSAria III flow cytometer, a high-speed fixed-alignment benchtop cell sorter. This document includes starting up the system, setting up the stream, checking cytometer performance with Cytometer Setup and Tracking (CS&T) beads, sorting, cleaning, and shutting down the system. 

The procedures apply to users operating the BD FACSAria III in room J.2.04, level 2 of WIMR. All personnel require training prior to independent operation of the instrument. Training is conducted by a trained operator or the scientific platform manager (if appropriate) with competency demonstration necessary before authorisation of access. Competency is assessed via demonstration of independent instrument operation, in conjunction with verbal explanation of all aspects of operation of the instrument and troubleshooting common and simulated faults. All instrument operation is to be conducted by trained operators.

Hours of operation and emergency contacts

The following hours of operation are valid from July 2nd 2018 unless otherwise updated.

Note: internal phones require a '0' to dial an outside line. Omit the listed '0' if calling from outside of WIMR. 

Reception / business hours

8.30am to 5.00pm

Monday to Friday

Advanced Specialist on 0-8627 1820

Scientific Platforms Manager on 0-8627 3210

For POLICE, AMBULANCE or FIRE emergencies contact 0-000

Outside of business hours

After hours contact the WIMR Emergency Management System contact on 0-0467 818 730

For access or security related issue contact TRICORP security on 0-1300 456 321

Research hours - WIMR

6.00am to midnight

Monday to Friday

8.00am to 8.00pm

Weekends and public holidays

Research hours - non-WIMR

7.00am to 9.00pm

Monday to Friday

8.00am to 8.00pm

Weekends and public holidays

Training and Competency Requirements

All users must have completed training.

Training is conducted via an initial theoretical introduction to the components and safety aspects of the instrument and laboratory followed by multiple sessions with a trained operator until competency in all aspects of safety and operation is demonstrated via independent operation, including dealing with emergency situations and performing all applicable tasks, and the skills necessary to describe functional components used while independently operating the instrument.

List of hazards and risk controls as per risk assessment

Associated risk assessment reference: WIMR-RA-OP-WS-504.02

Procedure

Definitions

In these procedures the following terms have the meaning set out below

1. BD – Beckton Dickinson
2. DI water – Deionized water
3. EtOH – Ethanol
4. Sheath solution – saline, 0.9% NaCl
5. SIP sheath - outer SIP steel cover
6. BSC – Biological safety cabinet (class 2 or above)
7. CST Beads – Cytometer Setup and Tracking Beads
8. SIP - Sample injection probe
9. FACS - fluorescence activated cell sorters 

List of resources 

(including personal protective clothing, chemicals and equipment needed)

• FACSAriaIII system
• Biosafety Safety Cabinet
• Sonicator
• PPE including gloves, long sleeve gowns, P2 respirator, safety glasses, enclosed shoes
• Bleach (12.5%) (Diluted 1 in 10 final)
• Decon 90 (concentrated surface decontaminant) (Diluted 1 in 20 final)
• Ethanol (70% w/w)
• Trigene (Diluted 1:50) or F10SC (Diluted 1:250) if Trigene isn’t available.

Biosafety considerations

• Samples can only be run on the fluorescence activated cell sorters occur after the approval of an associated project in PPMS.
• 100mL of 12.5% sodium hypochlorite (undiluted from the provided containers) must be added to the emptied waste containers at the start of the sorting session.
• All biological samples require filtering prior to acquisition on the FACS to prevent aerosol generation issues arising from clogs. 
  • Fixed samples may be filtered immediately before acquisition using the WRHFlow provided 50um filters and plastic tweezers. 
  • Unfixed samples may be filtered before acquisition using the WRHFlow provided 50um filters or the tube cap filters and plastic tweezers in a biological safety cabinet.

Procedure (Step by step instructions for undertaking the task)

PPE including gloves, long sleeve gowns, safety glasses, enclosed shoes must be worn before operating the instrument.

PPE including gloves, long sleeve gowns, safety glasses, enclosed shoes and a P2 respirator must be worn before operating the instrument in any event where there may be aerosol generation.

Samples that are to be acquired on the instrument are required to be transported to the flow cytometry labs in accordance with PC2 sample transport requirements.

1. Starting up the FACSAria III system
2. Turn on the CAS hood, chiller and aerosol management system (set at 20% - normal evacuation) (AMS).
3. Turn on the main switch to the ARIAIII.
4. Log into the computer as administrator
5. Log into the PPMS account as operator
6. Start BD FACSDiva Software.
7. Turn on the high-pressure gas and the cooling unit.
8. Relieve any pressure from the tanks by venting using the venting valve.
9. Fill sheath tank to weld mark (~10cm from top of tank) and reseal sheath tank.
10. Empty waste container slowly into the sink under running water. Put 100ml of bleach into the waste tank.
11. Reconnect waste container.
12. Check the filter attached to the sheath and at the front of the fluidic cart for air bubbles. Bleed out any air bubbles if required.
13. Allow ARIAIII to warm up for at least 30 min.
14. Ensure the computer is on and then log in to the administrator account. Username is “administrator”. Password is given to approved users for the account administrator.
15. BD FACSDiva should launch automatically. Once logged in to BD FACSDiva, the software will detect and connect to the instrument. If it does not do it automatically, click Instrument in the main menu and connect the instrument manually.
16. Sonicate the appropriate nozzle. Sonicate in DI water for 5 minutes in a 5mL FACS tube. Ensure sonicator has sufficient liquid in it to conduct cleaning power to the nozzle.
17. If needed - unlock the waste nozzle from the flow cell. Place the waste nozzle into an empty slot on the nozzle storage block.
18. Use a Kimwipe to remove any residue liquid on the nozzle to be installed if necessary.
19. Clean any salt residues around the nozzle assembly.
20. Insert sonicated nozzle with the O-ring facing upwards. Lock the nozzle.

1. Setting up the stream
2. If nozzle was changed, also change nozzle size settings in the software.
3. Open the sort block door, click the stream start/stop button to turn on the stream.
4. Check the stream video feed is not impeded by any liquid/debris on the lens. If this is the case it can be cleaned with a lens cleaning tip.
5. Check the stream angle. If the stream flows away from the centre of the waste catch, loosen the screws on both sides of the collection assembly and rotate the sort block to adjust.
6. Close the sort block and thumb screw it shut.
7. Inspect the stream for stable droplet generation, symmetrical droplets and similar breakoff position to the last time the stream was started. Click the sweet spot button and inspect the stream. Allow stream to stabilise. Adjust frequency and amplitude as needed with sweet spot off, updating values if necessary.
8. Do not continue the sort with an unstable stream.
9. Carefully check the instrument for wet areas indicating any leaks in the tubing or failing valves.

1. Checking Cytometer Performance with CS&T beads in DIVA.
2. Uncheck sweet spot. Select Cytometer > CS&T.
3. Select performance check. Ensure correct bead lot CST beads are loaded onto the instrument and click run.
4. If the beads fail, inspect the report and determine whether the stream and sample are as expected. If CS&T continues to fail call BD service.
5. If QC passes, view report to check any warnings.
6. Quit the CS&T window. Click use CS&T settings in Diva.

1. Checking drop delay (the time between the cytometer seeing an event and the event going into the last connected droplet) with Accudrop beads
2. Prepare Accudrop beads by adding 1 drop of beads into ~250ul of saline in a 5mL FACS tube. Diluted beads are OK to use within ~1 week.
3. Open the Accudrop experiment template. Bring out sort window from global worksheet.
4. Load Accudrop beads. Click sort, then click cancel on the dialog that appears to keep the waste drawer closed.
5. Turn on high voltage plates.
6. Make sure sweet spot is on. Using Accudrop beads, toggle optical filter on and adjust the Accudrop laser focus to obtain the brightest bead spot on the side stream.
7. Adjust drop delay to give the cleanest side streams by incremental increases/decreases.
8. Check, adjust if necessary and type in drop breakoff and gap values. Drop delay is now set.
9. Unload Accudrop beads.
10. Turn on test sort. Check if the side streams are clean and directed to indicated marks on the screen (i.e. going into the collection tubes intended to be used for the sort). If the side stream is slightly out of the centre of the collection tube, use the slider controls to adjust the side stream voltage. Turn off test sorts and high voltage plates.

1. Sorting
2. Ensure any disinfection solutions (1% bleach, 70% EtOH, 5% Decon, 1:50 Trigene, and any others necessary) are prepared before starting the sorter.
3. Create a new experiment or duplicate an existing one. Setup experiment using controls provided by the user. Filter controls samples before acquiring on a cytometer if necessary.
4. Install collection tubes into tube holder and slide into the sort block.
5. Filter the sample if needed
6. Load the filtered sample onto machine. Initially run the sample with a low flow rate in the acquisition window and record an appropriate number of events. Draw or adjust gates if needed.

• . Confirm approval for sorting strategy.

1. Input the population being sorted in the Sort Layout window. If doing a 4-way sort, use outer tubes for the lower frequency populations with the higher frequency populations in the middle two tubes. This is to reduce the chance of larger populations contaminating the smaller populations. There are some caveats to this, i.e. number of cells to be sorted, cell size, and side stream stability need to be also considered. Consider appropriate sort mask settings to be used.
2. Click Sort and turn on agitation to an appropriate value. Adjust flow rate and side streams if necessary. Threshold rate should not exceed approximately ¼ of the frequency to ensure events are spaced out along the stream. Monitor sort efficiency and adjust parameters if necessary.
3. Avoid running the sample dry to prevent bubbles in the system. If it’s a precious sample add more saline close to the end. When the sample level is low, reduce or halt agitation of the sample to minimise the chance of drawing in air. Monitor sample continuously and stop the sample before it is at a critically low level to prevent drawing air into the sample line.

1. Purity Check
2. Choose Instrument > Cleaning Modes > Sample Line Back flush. Click Start at prompt a few times to clean the valve, and wait for some time (~10-30sec) to clean the sample line, click Stop.
3. Load a tube of DI water and change flow rate to 11. Clean the tubing for at least 1 minute or until no sample looking events are displayed. Unload water tube and change flow rate to 2-4.
4. Dilute ~5ml of sorted sample into 100ml of diluent (4 drops of saline) in a new tube. Load the purity check tube and record sample for approximately 30-60 seconds.
5. Between running purity check tubes, perform sample line back flush.

1. Data export
2. Export FCS files & batch analyse samples when appropriate into default locations. Move the batch analysis into the FCS files folder ready for syncing or user transfer as setup for data management.

1. Daily shutdown of instrument
2. Load 10% bleach. Change flow rate to 11 and run for 5 min.
3. Repeat clean with 5% Decon 90.
4. Repeat clean with DI water.
5. Stop stream. Load 70% EtOH and initiate clean flow cell command. Repeat.
6. Remove nozzle and nozzle clip and replace to default locations.
7. Spray the sort chamber and other appropriate internal chambers specific to the instrument with 1:50 TRIGENE and wipe clean. Spray again with 1:50 TRIGENE and allow a contact time of at least 10 minutes.
8. Wipe the areas and follow with spraying and wiping clean with 70% ethanol
9. Spray the exposed appropriate external surfaces specific to the instrument with 1:50 TRIGENE and wipe clean. Spray again with 1:50 TRIGENE and allow a contact time of at least 10 minutes.
10. Wipe the areas sprayed and follow with spraying and wiping clean with 70% ethanol
11. System is now ready to be shut down. Turn off instrument, gas, chiller and hood afterwards. Log off computer.
12. Relieve any pressure from the tanks by venting using the venting valve.

Initial instrument startup, initialisation, setup & quality control

1. If a PPMS booking is made before 8:00am, WRHFlow will endeavour to have the instrument warmed up and ready to use by 9:00am. All instruments are booked from 8:00am until 9:00am for quality control procedures. However if the instrument is not on, turn on the flow cytometer by pressing the green power on button and allowing the lasers to warm up for 30 minutes.
2. Log on to the instrument computer with your WIMR active directory account.
3. Instrument acquisition software (BD FACSDiva) will automatically launch along with an internet browser by default showing the instrument user log and the booking schedule. 
4. Once Windows has loaded and FACSDiva has started, log in to FACSDiva with your group account from the drop down list. The password is the same as the group name in small caps.
   
5. If a CST mismatch dialog appears always click ‘Use CST Settings’
   
6. Check if CS&T (quality control) has been performed for the day on the user log that is displayed on the internet browser. Skip the below step if it has already been completed for the day.
   
   
   1. If CS&T hasn’t been completed, prime the instrument (LSRII/Fortessa/Symphony) and then run H20 for 3 minutes or perform a fluidics startup for the CantoII by clicking
      ‘Menu’ > ‘Cytometer’ > ‘Perform fluidics startup’.
   2. Once complete click
      ‘Menu’ > ‘Cytometer’ > ‘CS&T’. Select 'Check Performance', and ensure the correct bead lot number is selected. Each instrument will have a bead lot number displayed on the front of the instrument. Load CS&T beads from the WRHFlow fridge (normally premade, but can be made with 1 drop from the stock bottle in the flow fridge with 300uL saline in a 5mLs FACS tube – refer to MSDS) and click the run button. For the LSRII/Fortessa/Symphony, run beads on low with the dial 5 full rotations from either end. Take note of CST result. If successful, the instrument is ready to use, and the CS&T window can be closed. If CS&T fails, note the error and report it to flow staff before using.
      
7. If you are a new user create a folder per user under your group login using the following syntax ‘FirstnameLastnameinitial’. For example, Elizabeth Potter would create a folder called ElizabethP.
   
   

   Creating a new experiment

8. Create a new experiment under your folder, label the experiment using the following syntax ‘YYYYMMDD Descriptive_Name’.
9. Click ‘Cytometer Settings’.
10. Check the FSC-W, FSC-H, SSC-W, SSC-H parameters for recording in the Inspector tab as below to allow identification of doublets.
    
    
    Set up, create and calculate compensation for multicolour experiments
11. Click ‘Menu’ > ‘Experiment’ > ‘Compensation’ > ‘Create compensation controls’.
    
12. Leave the compensation tube labels as ‘Generic’ (unless you are using multiple panels in the same experiment as FACSDiva can calculate multiple compensation matrixes and match them with listed fluorochromes, WRHFlow recommends 1 experiment:1 fluorescent panel to avoid confusion). Ensure you have ‘Include separate unstained control tube’ checked (even if you are not using a universal control - as this provides a template displaying all detectors and can be removed later). 
    
13. Delete unused parameters from the list of fluorophores to be compensated.
14. Click OK when complete.
15. In the newly created compensation control folder, activate the ‘Unstained Control’ (as it has a useful default specific workspace that shows all instrument parameters).
    
    
16. Acquire any 1 of the compensation controls on low. Adjust FSC and SSC voltages to bring populations onto scale (take note of the FSC & SSC voltages needed to bring bead/cells on scale)
17. Sequentially acquire (*but do not record*) all the compensation controls, while the unstained tube is activated, ensuring for each channel that:
    1. Fluorescence signal is not off scale, ideally less than 2x10⁵ – if off scale reduce the parameter voltage from the cytometer settings window (shown below)
       
    2. Compensation control signals are higher than a fully stained sample – as if this isn't the case will result in an incorrect compensation matrix
18. After the above steps, your voltages should now be set. There are other methods to determine optimal detector voltages. Below is the BD method for determining the minimal (and not optimal) voltages that should be used. 
    1. • Acquire unlabelled cells. Adjust FSC and SSC voltages as needed to bring populations onto scale when looking at a FSC/SSC bivariate plot (take note of the voltages needed for cells).
       • Move the already drawn gate, P1, around the cells of interest. Right click a bi-variate plot and click ‘show population statistics’. Edit the statistics display to show the rSD for each fluorescent parameter used (see figure below). Record enough cells to determine the rSD of P1 for all parameters that will be used. You need to record a small file in order to accurately determine the rSD of the unlabelled cells.
         
       • Adjust PMT voltage to bring the rSD of the unlabelled cells to 2.5x the value of the rSD of electronic noise (that can be determined from the instrument configuration sheet) that accompanies each cytometer. You need to record a small file in order to accurately determine the rSD of the unlabelled cells each time.
       • Take note of these PMT voltages. These are the minimum PMT voltage that should be used for each detector for optimal detection of dim epitopes.
       • At this point you now have the minimum voltage for each detector needed to separate dim from negative signal - this voltage is not the voltage you should, but provides you with information on the lower bound of the detector voltage.This is the BD method for users wanting to know the minimal voltage that should be used (the voltage used should be higher than the minimal voltage).
19. Now that your voltages are set, you need to determine whether you are using a universal negative or not for compensation. If you are, then you can now record compensation controls - skip to step 20. Otherwise
    1. Click ‘Menu’ > ‘Experiment’ > ‘Compensation’ > ‘Modify compensation controls’. If you are using samples to compensate that have different levels of autofluorescence, we cannot use a universal negative. At this point, before you record the compensation controls uncheck the ‘Include separate unstained control tube/well' option.
       
       Bring up the compensation control window.
       
       Uncheck the universal negative option.
20. Now we will acquire the samples that will be used to calculate compensation. 
21. Activate the correct tube by clicking the arrow to the left of the tube. 
22. It is okay to change FSC and SSC detector voltages to those noted as you acquire different particles for your compensation controls.
23. Increase the events to record above the default number of 5000 events. We recommend at least 40000 events for a 6 colour experiment.
24. Gate the positive beads of interest for each fluorochrome as P2 as you acquire and record all the fluorescent compensation controls. If not using a universal negative (After recording each compensation control, create the positive and negative gate, by drawing a new P3 gate, as the software looks for a positive signal in the P2 gate and a negative signal in the P3 gate (see below)).
    
25. Calculate compensation by clicking
    ‘Menu’ > ‘Experiment’ > ‘Compensation’ > ‘Calculate compensation’, then apply link and save
    
26. You can leave the default compensation name or you can give the compensation settings a name following the syntax ‘YYYYMMDD_Descriptive_Name_Comp’
    

    Acquiring your samples

27. Create a new specimen in the same experiment and name relevant tubes with sample and control identifying information, additionally this information can be incorporated as metadata that is useful in offline analysis using the experiment layout menu item from the Experiment menu.
28. Modify the experiment layout to add all the names of the antibodies used for staining to the channel names. Modify the experiment layout to ensure you are recording and storing the correct number and set of events.
    
    
29. Activate the global worksheet (top left bottom on the worksheet toolbar).
    
30. Create an approximate gating strategy in the worksheet by drawing some initial plots to view data, the following are recommended to be shown if applicable with/out gating:
    1. Cells of interest (FSC-A x SSC-A)
    2. Single cell gate (FSC-A x FSC-H)
    3. Live dead gate (removal of dead cells)
    4. Fluorophores of interest (fluorophores can be displayed on a bi-variate plot).
    5. Time plot (TIME x last fluorescent parameter used)
       
31. Ensure that all fluorescent parameters are displayed in Log, and have bi-exponential display turned on from the Inspector window (unless linear signals are needed).
    
32. Activate the tube in the browser by ensuring there is a green arrow next to the tube you are to acquire and record data from.
33. Load your first sample to be acquired onto the instrument SIP. The CantoII and LSRII/Fortessa/Symphony have different setups.
    1. CantoII – move the sample adapter arm to the left, load tube, release sample adapter arm, then click acquire sample - be careful of a pinch hazard from the sample arm. 
    2. Fortessa/LSRII/Symphony – move the sample arm, remove the existing tube (if applicable, from the SIP), SIP can be washed by acquiring H20 between samples to minimise cross contamination, load the next tube and move the sample arm back under the tube.
34. Acquire data on low initially to adjust the FSC / SSC detector voltages to those noted for cells.
35. Now you can run and record your samples with an appropriate flow rate (the instrument electronics can handle ~20000 events per second). It is recommended that due to baseline restore the same flow rate is used for all test samples to be acquired. 
36. Acquire and record data from each of your samples.
37. Unless your data should not be synced to the WIMR server, export the data as FCS3.0 files in linear format to D:\BDExport\FCS (as this folder is automatically synced to the WIMR server). If you would like to copy your data elsewhere copy it from this location. Your export folder can be easily accessed from the quicklaunch toolbar. Your data should be able to be accessed from the Scientific Platforms networked drive within WIMR, and via a username/password for non-WIMR users if desired (email wrhflow@sydney.edu.au to setup).
    
    
    
    

    Cleaning the instrument

38. Clean the fluidics on the instrument by running the machine with 3 min 10% bleach, 3 min 5% decon & 3 min H2O on high.
39. Close the software.
40. Check to see whether you are the last user for the day. If you are the last user for the day turn off the instrument. For CantoII, if you are the last user of the week perform a fluidics shutdown.
41. Log out of the instrument PC, complete the user survey.
    
42. Empty the flow cytometer waste container. Add 100ml of bleach into waste tank and refill the sheath. Bleached waste is poured down the sink gently under a running tap, minimising splashing. All other laboratory biohazardous waste is to be removed via the biohazardous waste disposal bins located in the lab.
43. Clean up and decontaminate the area with 70% ethanol.
44. Turn off the instrument using the green power button if you are the last instrument for the day.
    

    Data management

• Under no circumstances should there be any patient/person identiable data recorded on the instruments as the data is not privately held and is accessible to all researchers. 
• User data is deleted after 7 days off the instrument computers, it is the responsibility of the user to have transferred data to a secure location.
• WRHFlow recommends exporting data from BD FacsDiva as FCS3.0 files in linear format to D:\BDExport\FCS. This folder is automatically synced to the WIMR Scientific Platforms Drive that all WIMR active directory users can access. For the sync to work properly your experiment must be in a folder with "FirstnameLastinitial" syntax. Please note this folder is accessible to all researchers.
• If you would like your data accessible from outside of WIMR, a user specific share can be setup. This works by creating a copy of a specified folder on university provisioned storage that is shared with the user. This requires an email to wrhflow@sydney.edu.au and it is the users responsibility that all ethics approvals are complied with. 

Application Settings

Application settings are useful in ensuring longitudinal studies (studies measured over multiple days/weeks/months) maintain consistent MFIs over the course of the experiment. We have successfully implemented and assisted many users in creating and applying application settings to their experiments.

The following workflow can be used to successfully save and apply application settings. Please note this is only one workflow to save and apply application settings. There are several variations that could also work (and some that might not). We are covering 1 specific workflow today. The 2 key points to remember are, 1, that you check that voltages have updated to ensure successful application of application settings, and 2, to record data for beads (CST, with bead lot recorded are good for this, or other long term QC beads may be used) before and following your experiment, every day you collect data. Running these beads allow continuation of targeting similar MFIs, even if the instrument undergoes major change, or if the baseline settings expire for application settings.

The below listed protocol is for creating a new experiment with N compensated parameters. It assumes you have 1 panel with N flurochromes, and will setup and apply a compensation matrix to your data when starting. The assumption is that you have compensation controls for all parameters. This protocol creates application settings that are used on subsequent days to ensure consistent MFI’s. We will be documenting a different protocol for setting up application settings with an exported experiment template as there are slight variations

Experiment Setup

• Make a new specimen and tubes as necessary.
• Menu > ‘Experiment’ > ‘Experiment Layout’
• Label all parameters, setup acquisition parameters, storage gates, event numbers, and input keywords.
• Rename specimens / tubes as needed.
• Setup workspace layout as necessary. A good workspace observes all parameters in strategic bivariate plots, and monitors the acquisition over time by drawing a parameter using last laser on the Y axis with Time on the X axis. A good experiment will also have a tube where the user can record beads (QC beads, 8 peak, CAB beads, etc.) both as the first tube and the last tube in the experiment, ensuring that there was no aberrant instrument occurrences during the acquisition and also to allow adjusting of PMT voltages in the event of instrument component replacement or instrument baseline update.
• Load your first sample to be acquired and recorded. Activate the tube in the browser. Acquire data, and quickly adjust the FSC / SSC detector voltages to those noted for cells. Record your first samples.
• Right click the top-level cytometer settings and click ‘Application Settings’ > ‘Save’.
• Give the application settings a name following the syntax ‘YYYYMMDD_Descriptive_Name_AppSettings’.
• Acquire and record data for each sample. - including reference beads.
• Export data.
• Note down all experiment details.

 Restoring application settings (from the original experiment, not from experiment template)

• Open the original experiment (for this protocol, do not open an experiment that has already had application settings restored, only open the original experiment).
• Right click experiment > ‘Duplicate experiment without data’.
• In the newly created duplicated experiment, right click the duplicated experiments top-level cytometer settings ‘Cytometer Settings’ > ‘Application Settings’ > ‘Apply’ > Apply saved application settings
• As the copied experiment will have a compensation matrix that is non-zero from when the experiment was setup, click ‘keep the compensation values’ (you can re-record compensation control tubes and recalculate the compensation matrix if desired).
• If a dialog appears asking to ‘Confirm Cytometer Changes’ – in regards to FSC Area Scaling values have changed – overwrite them by clicking YES.
• If you are not recording new compensation controls, proceed to record samples (adjusting FSC and SSC detector voltages to those noted if necessary).
• If you are recording new compensation controls (recommended), right click the compensation tubes ‘Cytometer Settings’ > ‘Application Settings’ > ‘Apply’ > Apply saved application settings. Record compensations (only adjusting FSC and SSC detector voltages (as noted) to bring populations onto scale, then calculate compensation. Clicking apply and save.
• Check the voltages have been slightly updated.
• Proceed to record samples (adjusting FSC and SSC detector voltages to those noted if necessary)

Notes

• Remember there is a range of optimal PMT voltages that will provide similar resolution between dim and negative cells.
• PMT voltages that are too high will not affect the resolution of dim and negative cells, but can unnecessarily cause spreading of the negative population.
• Any bright populations with very large CVs (very broad distributions) must have PMT voltages lowered so that the brightest events or MFIs remain within the linear range of the detector.
• Remember your compensation controls should be representative of your samples, ensuring they are as bright or brighter than your sample (but not log folds brighter preferentially).
• Remember to leave enough room above the positive signal, and in the linear range of the detector, to allow for increases in fluorescence of certain epitopes.
• We recommend your samples are filtered beforehand.

Additional Information

Contamination testing sample collection – For contamination testing samples from the tanks, stream and sample tubing are collected for downstream contamination testing. In addition to the above procedures the following need to be undertaken when obtaining these samples.

• Sheath tank: Ensure tank pressure is relieved prior to opening the sheath tank for sample collection. Only collect tank samples using a sterile collecting means into a sterile 5mL FACS tube.
• Stream: Collect sample using the test sort mode on the instrument into a collection tube.
• Sample line: Using a 5mL falcon tube that has 4mL of sterile saline in it, load and unload a sample 3 times to sample the sample line.

Routine instrument / lab maintenance – For the laboratory to function efficiently lab/instrument maintenance will include

• Stocking up saline bottles, H2O bottles, gowns, P2 respirators, sterile pipettes, tubes, markers, stationary, reagents and other unlisted as required in the laboratory work flow.
• Stocking up cleaning solutions including bleach, Decon 90, 1:50 TRIGENE & ethanol.
• Making up cleaning solutions daily
• Disconnecting the depressurised sheath tank for autoclaving, sheath filter installation, waste tank filter replacement, instrument cleaning and performance checking.
• Autoclaving empty wash bottles holding bleach, ethanol, H20 and refilling under sterile conditions.
• Reconnecting fluidic lines as per instrument setup.

Instrument decontamination prior to service – The instrument should have been decontaminated prior to access for service.

• Spray the sort chamber and other appropriate internal chambers specific to the instrument with 1:50 TRIGENE and wipe clean. Spray again with 1:50 TRIGENE and allow a contact time of at least 10 minutes.
• Wipe the areas and follow with spraying and wiping clean with 70% ethanol.
• Spray the exposed appropriate external surfaces specific to the instrument with 1:50 TRIGENE and wipe clean. Spray again with 1:50 TRIGENE and allow a contact time of at least 10 minutes.
• Wipe the areas sprayed and follow with spraying and wiping clean with 70% ethanol.
• No further biological samples are to be run on the instrument until maintenance is carried out.

Emergency procedures

All emergencies need to be reported to the emergency contacts listed above. Specific chemical exposure procedures are below.

Bleach

Eye exposure:

1. Wash out immediately with saline from the emergency eye wash bottle.
2. Ensure complete irrigation of the eye for at least 15 minutes by keeping eyelids apart and away from eye and moving the eyelids by occasionally lifting the upper and lower lids.
3. Seek medical attention immediately.
4. Removal of contact lenses should only be done by medical personnel.
5. Notify emergency contacts

Skin exposure:

1. Remove all contaminated clothing immediately.
2. Flush affected area under running water.
3. Seek medical attention if necessary.
4. Notify emergency contacts

Emergency power off button activation if an electrical, or laser hazard is present

1. Note that the emergency power shutdown will only isolate the red powerpoints. 

2. If there is an electrical hazard and the instrument is plugged into a power point that is not on the uninterruptible power supply (blue power point), pressing the emergency power off button will stop electricity to the instrument. Note that if an electrical device is plugged into the blue power point it will continue to be live. In this event leave the room, ensure a message is placed on the door alerting others and seek assistance.

3. In an electrical emergency press the red power shutdown button in the room.
4. Leave the room and place signage that the room is not to be entered due to a major electrical problem.
5. Notify emergency contacts

Emergency high pressure gas shutdown

If there is an issue with the high-pressure gas, the gas supply can be shut down by closing the valve on the wall and or the secondary shut off valve located in the main corridor outside of room J2.04.

Biohazard spill 

1. Obtain biohazard spill kit - located in all flow cytometry labs - see appendix for location.
2. Remove contaminated PPE and place in biohazard waste bag; and
3. Put on waterproof gown, gloves and mask with eye protection before proceeding.
4. Sprinkle CliniSorb powder over the spill; or
5. Cover the area of the spill with disposable cloth soaked in freshly prepared disinfectant suitable for the material in use (1% sodium hypochlorite - 100mL of 12.5% bleach in 1L bottle, or 1%Virkon - 2 tablets in 1L bottle); and
6. Leave it for at least 10 minutes.
7. Apply second cloth soaked in disinfectant on top of the first;
8. Use a third cloth (dry) to pick up and transfer the cloths to waste bags. If CliniSorb powder was used, collect the spilt material and Clinisorb powder mixture using spatula/scraper. Discard to waste bags;
9. Decontaminate the area thoroughly with suitable disinfectant;
10. Dispose of all waste including gloves in waste bags;
11. Notify emergency contact who can for DNIR spill, double bag the waste in autoclavable bags and loosely seal with autoclave tape for removal for autoclaving.

Chemical spill 

1. If you are in doubt of your ability to clean a chemical spill safely then evacuate the area and seek help. It is better to ask for help if you are not sure of how to clean it up properly.
2. If there is risk to the rest of the building contact the emergency contacts to initiate building evacuation.
3. Review the SDS and make sure you understand the hazardous properties of the spilled material before you attempt to clean it up.
4. First aid is always the top priority. If a hazardous material is spilt on yourself, remove any potentially contaminated clothing immediately and use the emergency shower. If a chemical spill has entered your eyes flush for at least 15 minutes at the eyewash station. Seek appropriate medical treatment.
5. Determine whether it is a major or minor spill. A major spill is one that involves a large amount of hazardous material, poses a risk of fire/explosion, a respiratory hazard exists, or when dealing with unknown chemical spills. Seek assistance from emergency contacts for major spills. 
6. For minor spills alert lab manager, safety officer and others in the lab and cordon off the affected area. Isolate the spill.
7. Retrieve the spill kit (there are also adequate instructions in the chemical spill kit). Located in the shared prep lab J2.06 - see appendix for location. Stop and think about your plan to clean the spill. Remove the gloves and goggles and from the kit, put them and all appropriate PPE on before approaching the spill.
8. Identify the spill - there is a pH strip tester that can be used, also can refer to SDS, labels, supervisors. 
9. Select the agent to clean up the spill.
   1. For acid spills: Spill-X-A® acid neutraliser
   2. For caustic spills: Spill-X-C ® caustic neutraliser
   3. For solvent spills: Spill-X-S ® solvent adsorbent
   4. For formaldehyde spills: Spill-X-FP ® formaldehyde
      polymeriser
   5. In some instances combinations of adsorbents many be necessary. 
10. Encircle spill, cover with adsorbent
11. Mix adsorbent into spill
12. Using the absorbent pads from the spill kit, carefully wipe up the
    spilled liquid, again working from the outside in.
13. Beware of any neutralising reactions
14. Collect powdery residue using spatula, deposit in waste bags
15. Place all waste materials in a plastic bag. Once the spill has been
    fully cleaned, place the waste bag with in the fume hood temporarily.
16. Label and dispose of waste according to building policies. 
17. Remove PPE and thoroughly wash hands.
18. Report the spill using the buildings incident notification form.

1. Emergency shutdown for stream failure:

If a stream failure occurs, the AriaIII may detect this and shut off the stream and sample automatically. If this occurs during sample being put through the instrument, the instrument must be given time for any potential aerosols to settle or be removed via the biological safety cabinet before accessing the sort block.

If the instrument does not automatically shut down the stream upon a stream failure, the stream sample and stream need to be stopped via the software or via the red emergency button located on the instrument. Then after a period of 15 minutes, and with the appropriate PPE (gloves, gown, enclosed shoes and safety respirator), the nozzle may be removed if necessary, decontaminated and sonicated if necessary to remove the clog and reinstalled if needed. If the stream failure was due to a clogged sample, the sample may need to be re-filtered or a larger nozzle used. The primary objective is to prevent potentially biohazardous aerosols being exposed outside of the biological safety cabinet. The procedure is outline below.

1. Ensure BSC is on
2. Ensure PPE is in use
1. Gloves, gown, safety glasses & P2 respirator
3. Stop the stream if the instrument hasn’t automatically. The stream can be stopped via the software or the emergency red button on the instrument
4. Minimise BSC turbulence
5. Keep sort chamber closed and sample cover closed to contain aerosols
6. Allow 15 minutes for the BSC to evacuate any aerosols
7. Remove collection tubes and sample aseptically

1. Spray the sort chamber, sort collection block and other appropriate internal chambers specific to the instrument with 1:50 TRIGENE and wipe clean. Spray again with 1:50 TRIGENE and allow a contact time of at least 10 minutes.

1. Determine cause of sort failure if possible.
2. Remove nozzle and sonicate if necessary.
3. Restart the stream if possible.
4. If nozzle clog occurred – refilter sample or use larger nozzle before loading sample.
5. Log the failure in the sort log.

1. Emergency shutdown if an electrical hazard is present

If there is an electrical hazard and the instrument is plugged into a power point that is not on the uninterruptible power supply (blue power point), pressing the emergency power off button will stop electricity to the instrument. Note if an electrical device is plugged into the blue power point it will continue to be live. In this event leave the room, ensure a message is placed on the door alerting others and seek assistance.

If there is an issue with the high-pressure gas, the gas supply can be shut down by closing the valve on the wall and or the secondary shut off valve in the corridor.

Associated Documents

This Standard Operating Procedure should be read in conjunction with:

1. Australian/New Zealand Standard - Safety in Laboratories Part 3: Microbiological safety and containment (AS/NZS 2243.3:2010)
2. Australian/New Zealand Standard - Management of clinical and related wastes (AS/NZS 3816:1998)
3. University of Sydney Safety Health & Wellbeing: Guideline for the Decontamination of Clinical/Biological Waste and Spill Management, http://sydney.edu.au/whs/guidelines/biosafety/decontamination_guidelines.shtml#2.2.1.
4. “A Guide to the WIMR Tissue Culture Facilities” – WIMR laboratory guideline

5. 
   

   BDFACS ARIA III User Manual – Scientific Platforms Network Drive
   1. Australian/New Zealand Standard - Safety in Laboratories Part 3: Microbiological safety and containment (AS/NZS 2243.3:2010)
   2. Australian/New Zealand Standard - Management of clinical and related wastes (AS/NZS 3816:1998)
   3. University of Sydney Safety Health & Wellbeing: Guideline for the Decontamination of Clinical/Biological Waste and Spill Management, http://sydney.edu.au/whs/guidelines/biosafety/decontamination_guidelines.shtml#2.2.1.
   4. “A Guide to the WIMR Tissue Culture Facilities” – WIMR laboratory guideline
   5. Holmes KL, Fontes B, Hogarth P, et al. International Society for the Advancement of Cytometry Cell Sorter Biosafety Standards. Cytometry Part A : the journal of the International Society for Analytical Cytology. 2014;85(5):434-453. doi:10.1002/cyto.a.22454.
6. MSDS - CST beads
7. WIMR – Management of Hazardous Materials Policy & Procedure
8. WIMR – Critical Risk Management Plan for biosafety
9. WIMR – Critical Risk Management for Waste Management
10. WIMR – Critical Risk Management Plan for chemical safety+
11. OGTR Guidelines - http://www.ogtr.gov.au/

Appendix

Chemical and biological spill kit location

Chemical Spill Kit Instructions

Biohazard Spill Procedure Poster

Health Risk Matrix used in risk assesment

Instrument diagrams depicting main components of ARIAIII