Date of revision and approval

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

Date of approval: TBD

Approval authority: Scientific Platforms Manager, Workplace

Date of revision and approval

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

Date of approval: 12 Dec 2018 

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

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Enquiries contact: weatmead.cytometry@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 located at the Westmead Research Hub. 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. It also includes safety considerations and emergency procedures. 

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. OGTR requirements for safe work in a PC2 laboratory apply.

Hours of operation and emergency contacts

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

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Advanced Specialist on 0-8627 1820Scientific Platforms 3216

Cytometry/Imaging/EM Manager on 0-8627 3210For 1820

Scientific platforms Director on 0-8627 3210

Email: westmead.cytometry@sydney.edu.au

For POLICE, AMBULANCE or FIRE emergencies contact 0-000

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Weekends, public holidays and shutdown periods. 

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 assisted sessions with a trained operator followed by independent operation until competency in all aspects of safety and operation are demonstrated via independent operation, including dealing with emergency situations and performing all applicable tasks. Description and purpose of functional components used while independently operating the instrument are also required. 

List of hazards and risk controls as per risk assessment

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 sorting 

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 1:50100) 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.
• Sheath tank must be filled prior to stream startup to prevent cell sort failure midrun. 
• Biological samples can only be run on the fluorescence activated cell sorters occur after filtration. 

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.

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.

Starting up the FACSAria III system

1. Turn on the CAS BSC hood, chiller, high-pressure gas and aerosol management system (set at 20% - for operational velocity evacuation).
2. Turn on the main switch to the ARIAIII located on the left of the instrument. 
3. Log into Windows using provided credentials. 
4. Log into the PPMS account with provided credentials. 
5. BD FACSDiva should launch automatically as should Google Chrome (contains the sorting log).
6. Log 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.
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7. If a CST mismatch dialog appears always click ‘Use CST Settings’
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8. Open the BCS BSC cabinet.
9. Open the front cover of the AriaIII and prob prop it open.
10. Determine if the correct nozzle is in place in the flow cell.If it isn't .
    1. If the cleaning nozzle is in place perform a fluidics startup (as this indicates the fluidics was shutdown).
    2. If the incorrect nozzle is in place in the flow cell, sonicate the appropriate nozzle 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.
       1. If needed - unlock and remove the previous nozzle or waste nozzle from the flow cell. Place the waste nozzle into an empty slot on the nozzle storage block.
       2. Use a Kimwipe to remove any residue liquid on the sonicated nozzle to be installed if necessary.
       3. Clean any salt residues around the nozzle assembly area using ethanol. Wipe and dry clean. 
       4. Insert sonicated nozzle with the O-ring facing upwards. Lock the nozzle using the nozzle clip.
       5. Since the nozzle was changed, also change nozzle size settings in the software by setting the correct cytometer configuration - >Cytometer>Configurations>Select >Cytometer>View Configurations>Select correct nozzle>Applynozzle>Set Configuration. Open the sort block door, click the stream start/stop button to turn on the stream.
       6. 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.Check the stream angle.
    3. 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.
    4. Close the sort block and thumb screw it shut.
    If the nozzle wasn't changedcorrect nozzle is in place in the instrument, ensure the correct settings are instrument configuration is selected and applied in the software by looking at the window title bar. 
11. The ARIAIII flow cell cover should be open.
12. Open the sort chamber (we need to view whether the stream is aligned or not). 
13. Close the front cover of the BSC.
14. Be ready to change the waste catch position and then start the stream.
15. Change the waste catch position if necessary. 
16. Stop the stream. This starts air into the sheath tank which is used to minimise contamination of the sheath tank when refilling. Spray the sheath tank lid with ethanol.
17. Unscrew the sheath tank lid completely.Remove the air pressurising line.
18. Relieve all pressure from the tank by venting using using the venting valve, and breaking the air-pressure seal by pushing down on the lid. Ensure you keep fingers clear of the edges of the lid and remove it. It can be stored temporarily on the fluidics cart (keeping the inner face not touching anything).Connect the air pressurising line promptly.
19. Fill sheath tank to the weld mark (~10cm from top of tank).
20. Reseal sheath tank - only holding the screw mechanism to keep fingers clear of the lid. 
21. Empty waste container slowly into the sink under running water. Put 100ml of bleach into the waste tank.
22. Reconnect waste container.
23. Check the filter attached to the sheath and at the front of the fluidic cart for air bubbles. If needed, ethanol spray and then gently bleed out any air bubbles if required from the finger screw vent on the filter.  
24. Start the stream again. The stream should be aligned in the centre of the waste catch. 
25. Close the sort chamber doorTighten the vent. 
26. Be ready to change the waste catch position, loosen the screws on both sides of the collection assembly, and then start the stream.
27. Check the stream angle. If the stream flows away from the centre of the waste catch, rotate the sort block to adjust. Tighten screws.
28. Close the sort block and thumb screw it shut.
29. Stop the stream. 
30. Start the stream again. The stream should be aligned in the centre of the waste catch. If needed adjust. 
31. Open the front cover of the BSC hood and close the front of the ARIAIII lid. 
32. Close the front cover of the BSC.
33. Inspect the stream for stable droplet generation, symmetrical droplets and similar breakoff position to the last time the stream was started. Click Check for satellite droplet merging typically within 3 drops of breakoff. Click the sweet spot button and inspect the stream. 
34. Allow ARIAIII stream/lasers to warm / stabilise for at least 30 min.
35. Adjust frequency and amplitude as needed with sweet spot off, updating sweet spot values if necessary.
36. Carefully check the instrument for wet areas indicating any leaks in the tubing or failing valves.
37. Do not continue the sort with an unstable stream or leaks. Allow ARIAIII stream/lasers to warm / stabilise for at least 30 min.

Checking Cytometer Performance with CS&T beads in DIVA.

1. Uncheck sweet spot. Select Cytometer > CS&T.
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. Ensure tube is labelled appropriately. ) and click the run button. Take note of CST result. If successful, the instrument alignment is right and good 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.
   
3. Select performance check. Ensure correct bead lot CST beads are loaded onto the instrument (Correct bead lot is displayed on asign on the instruement) 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.

Checking drop delay (the time between the cytometer seeing an event and the event going into the last connected droplet) with Accudrop beads

1. Use prepared or if needed 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. Ensure tube is labelled appropriately. 
2. Open the Accudrop experiment template. Bring out sort window from global worksheet.
3. Load Accudrop beads. Click sort, then click cancel on the dialog that appears to keep the waste drawer closed.
4. Turn on high voltage plates.
5. Make sure sweet spot is on and the stream has been optimised. Using Accudrop beads, toggle optical filter on and adjust the Accudrop laser focus to obtain the brightest bead spot on the side stream.
6. Adjust drop delay to give the cleanest side streams by incremental increases/decreases.
7. You may need to increase / decrease sample flow rate. You may need to sweep the accudrop laser to intensify the accudrop beads visible on the camera. 
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.

Sorting

1. Ensure any disinfection solutions (1% bleach, 70% EtOH, 5% Decon, 1:50 Trigene, and any others necessary) are prepared before starting the sorter.
2. Create a new experiment or duplicate an existing one. Follow appropriate experiment creation workflow, i.e. syntax YYYYMMDD DescriptionSetup in a folder with the following syntax ‘FirstnameLastnameinitial’. Check the FSC-W, FSC-H, SSC-W, SSC-H parameters for recording in the Inspector tab as below to allow identification of doublets. Setup experiment using controls provided by the user. Filter controls biological samples before acquiring on a cytometer if necessary.
3. Install collection tubes into tube holder and slide into the sort block.
4. Filter the sample if needed
5. 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.
6. Acquire, record and sort data/cells from each of your samples.
7. Confirm approval for sorting strategy.
8. If needed create a new sort layout form the >Sort menu. 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. On the AriaIII, purity or 4way purity modes are commonly used and recommended. 
9. Install the correct and appropriate collection tubes in the sort collection chamber to collect sorted events. 
10. 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.
11. Ensure the 'Sort in Progress' signage is on the door. When the sort is finished this sign can be removed.
12. 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.

Purity Check

1. 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.
2. Load a full tube of DI water and change flow rate to The sort chamber can only be opened when there is no sample running. Ensure the waste draw is closed before opening sort chamber. Remove collection tube when sort is finished. 
3. Remove sample from instrument. 
4. 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 data should be able to be accessed from the Scientific Platforms networked drive within WIMR.
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Purity Check

1. 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.
2. Load a full 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.
3. Dilute ~5uL of sorted sample into 100uL of diluent (4 drops of saline) in a new tube. Ensure sample agitator is off. Load the purity check tube and record sample for approximately 30-60 seconds.
4. Between running purity check tubes, perform sample line back flush.

Data export

1. 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.

Daily shutdown of instrument

1. Load 10% bleach. Change flow rate to 11 and run for 5 min.
2. Repeat with 5% Decon 90.
3. Repeat with DI water.
4. Stop stream.
5. If the next sort booked on the instrument in that week uses the same nozzle
   1. Load a full tube of 70% EtOH and initiate clean flow cell command. 
6. If the next sort booked on the instrument in that week uses a different nozzle
   1. Remove current nozzle. 
   2. Replace with cleaning nozzle.
   3. Load a full tube of 70% EtOH and initiate clean flow cell command. 
   4. When complete - remove cleaning nozzle and remove nozzle clip.
7. If there is no other sort booked for the week. 
   1. Remove current nozzle. 
   2. Replace with cleaning nozzle.
   3. Initiate fluidics shutdown program from cytometer menu and follow prompts.
   4. Complete fluidics shutdown. 
   5. Remove cleaning nozzle and nozzle clip and replace to default locations.
8. Spray the sort chamber and other appropriate internal chambers specific to the instrument as well as exposed appropriate external surfaces with 70% w/v ethanol and wipe clean. Spray again with 70% w/v ethanol and allow a contact time of at least 10 minutes.
9. Wipe the areas and follow with spraying and wiping clean with 70% ethanol
10. System is now ready to be shut down. Turn off instrument, gas, chiller, AMO & BSC afterwards. Log off computer.
11. Relieve any pressure from the tanks by venting using the venting valve.
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12. If a CST mismatch dialog appears always click ‘Use CST Settings’
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13. 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.
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    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.
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14. 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.
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    Creating a new experiment

15. Create a new experiment under your folder, label the experiment using the following syntax ‘YYYYMMDD Descriptive_Name’.
16. Click ‘Cytometer Settings’.
Check the FSC-W, FSC-H, SSC-W, SSC-H parameters for recording in the Inspector tab as below to allow identification of doublets.
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Set up, create and calculate compensation for multicolour experiments
17. Click ‘Menu’ > ‘Experiment’ > ‘Compensation’ > ‘Create compensation controls’.
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18. 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). 
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19. Delete unused parameters from the list of fluorophores to be compensated.
20. Click OK when complete.
21. In the newly created compensation control folder, activate the ‘Unstained Control’ (as it has a useful default specific workspace that shows all instrument parameters).
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22. 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)
23. 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)
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    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
24. 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.
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       • 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).
25. 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.
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       Bring up the compensation control window.
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       Uncheck the universal negative option.
26. Now we will acquire the samples that will be used to calculate compensation. 
27. Activate the correct tube by clicking the arrow to the left of the tube. 
28. It is okay to change FSC and SSC detector voltages to those noted as you acquire different particles for your compensation controls.
29. Increase the events to record above the default number of 5000 events. We recommend at least 40000 events for a 6 colour experiment.
30. 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)).
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31. Calculate compensation by clicking
    ‘Menu’ > ‘Experiment’ > ‘Compensation’ > ‘Calculate compensation’, then apply link and save
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32. 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

33. 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.
34. 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.
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35. Activate the global worksheet (top left bottom on the worksheet toolbar).
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36. 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)
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37. 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).
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38. 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.
39. 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.
40. Acquire data on low initially to adjust the FSC / SSC detector voltages to those noted for cells.
41. 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. 
42. Acquire and record data from each of your samples.
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).
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Cleaning the instrument

43. Clean the fluidics on the instrument by running the machine with 3 min 10% bleach, 3 min 5% decon & 3 min H2O on high.
44. Close the software.
45. 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.
46. Log out of the instrument PC, complete the user survey.
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47. 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.
48. Clean up and decontaminate the area with 70% ethanol.
49. Turn off the instrument using the green power button if you are the last instrument for the day
    1. nozzle clip and replace to default locations.
50. Spray the sort chamber and other appropriate internal chambers specific to the instrument as well as exposed appropriate external surfaces with 70% w/v ethanol and wipe clean. Spray again with 70% w/v ethanol and allow a contact time of at least 10 minutes.
51. Wipe the areas and follow with spraying and wiping clean with 70% ethanol
52. System is now ready to be shut down. Turn off instrument, gas, chiller, AMO & BSC afterwards. Log off computer.
53. Relieve any pressure from the tanks by venting using the venting valve

Notes

Detectors

• 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.

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@sydneywestmead.cytometry@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.

...

• 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..
• 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)

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.

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• 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, 1:50 100 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 autoclavable empty wash bottles holding bleach, ethanol, H20 for sterile H20 / saline and refilling under sterile conditions.
• Reconnecting fluidic lines as per instrument setup.

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• Spray the sort chamber and other appropriate internal chambers specific to the instrument with 1:50 100 TRIGENE and wipe clean. Spray again with 1:50 100 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 1:50 100 TRIGENE and wipe clean. Spray again with 1:50 100 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.

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• be run on the instrument until maintenance is carried out.

Emergency procedures

Emergency procedure folder can be found here in the sample prep lab:

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You can also access all these information via WIMR intranet:

https://intranet.westmeadinstitute.org.au/workingatwmi/HandS/Pages/Sarah%20Johnston.aspx

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

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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.

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 3 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 (provided)
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
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.

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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.

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1. Increase AMO to maximum flow rate to facilitate the evacuation of aerosols
2. Leave the room. 
3. Allow 3 minutes for the AMO & BSC to evacuate any aerosols
4. Remove collection tubes and sample aseptically.
5. Spray the sort chamber, sort collection block and other appropriate internal chambers specific to the instrument with 1:100 TRIGENE and wipe clean. Spray again with 1:100 TRIGENE and allow a contact time of at least 10 minutes.
6. Determine cause of sort failure if possible.
7. Remove nozzle and sonicate if necessary.
8. Restart the stream if possible.
9. If nozzle clog occurred – refilter sample or use larger nozzle before loading sample.
10. Log the failure in the sort log.

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
6. Australian/New Zealand Standard - Safety in Laboratories Part 3: Microbiological safety and containment (AS/NZS 2243.3:2010)
7. Australian/New Zealand Standard - Management of clinical and related wastes (AS/NZS 3816:1998)
8. 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.
9. “A Guide to the WIMR Tissue Culture Facilities” – WIMR laboratory guideline
10. 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.
11. MSDS - CST beads
12. WIMR – Management of Hazardous Materials Policy & Procedure
13. WIMR – Critical Risk Management Plan for biosafety
14. WIMR – Critical Risk Management for Waste Management
15. WIMR – Critical Risk Management Plan for chemical safety+
16. OGTR Guidelines - http://www.ogtr.gov.au/

Appendix

Chemical and biological spill kit location

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Chemical Spill Kit Instructions (located in J.2.06 - not Cell Sorter Lab)

Biohazard Spill Procedure Poster

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Health Risk Matrix used in risk assesment

Sort failure procedure for operators

Image AddedInstrument diagrams and room images depicting main components of ARIAIII system and room.

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