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

Functional unit: Flow Cytometry

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

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.

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 3216

Cytometry/Imaging/EM Manager on 0-8627 1820

Scientific platforms Director on 0-8627 3210

Email: westmead.cytometry@sydney.edu.au

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 / non-WIMR

6.00am to midnight

Monday to Friday

8.00am to 8.00pm

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 1:100) 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.
   
7. If a CST mismatch dialog appears always click ‘Use CST Settings’
   
8. Open the BSC cabinet.
9. Open the front cover of the AriaIII and prop it open.
10. Determine if the correct nozzle is in place in the flow cell.
    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 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>View Configurations>Select correct nozzle>Set Configuration. 
       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.
    3. If the correct nozzle is in place in the instrument, ensure the correct 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. Spray the sheath tank lid with ethanol.
15. Unscrew the sheath tank lid completely.
16. Relieve all pressure from the tank by venting 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).
17. Fill sheath tank to the weld mark (~10cm from top of tank).
18. Reseal sheath tank - only holding the screw mechanism to keep fingers clear of the lid. 
19. Empty waste container slowly into the sink under running water. Put 100ml of bleach into the waste tank.
20. Reconnect waste container.
21. 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. Tighten the vent. 
22. Be ready to change the waste catch position, loosen the screws on both sides of the collection assembly, and then start the stream.
23. Check the stream angle. If the stream flows away from the centre of the waste catch, rotate the sort block to adjust. Tighten screws.
24. Close the sort block and thumb screw it shut.
25. Stop the stream. 
26. Start the stream again. The stream should be aligned in the centre of the waste catch. If needed adjust. 
27. Open the front cover of the BSC hood and close the front of the ARIAIII lid. 
28. Close the front cover of the BSC.
29. Inspect the stream for stable droplet generation, symmetrical droplets and similar breakoff position to the last time the stream was started. Check for satellite droplet merging typically within 3 drops of breakoff. Click the sweet spot button and inspect the stream. 
30. Allow ARIAIII stream/lasers to warm / stabilise for at least 30 min.
31. Adjust frequency and amplitude as needed with sweet spot off, updating sweet spot values if necessary.
32. Carefully check the instrument for wet areas indicating any leaks in the tubing or failing valves.
33. Do not continue the sort with an unstable stream or leaks. 

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 biological samples before acquiring on a cytometer.
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.
13. 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. 
14. Remove sample from instrument. 
15. 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.
    
    

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

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 westmead.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.

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)

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, 1:100 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 for sterile H20 / saline 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 1:100 TRIGENE and wipe clean. Spray again with 1:100 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 1:100 TRIGENE and wipe clean. Spray again with 1:100 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.

Emergency procedures

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

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.

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.

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 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. Increase AMO to maximum flow rate to facilitate the evacuation of aerosols
7. Leave the room. 
8. Allow 3 minutes for the AMO & BSC to evacuate any aerosols
9. Remove collection tubes and sample aseptically.
10. 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.
11. Determine cause of sort failure if possible.
12. Remove nozzle and sonicate if necessary.
13. Restart the stream if possible.
14. If nozzle clog occurred – refilter sample or use larger nozzle before loading sample.
15. 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

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

Biohazard Spill Procedure Poster

Health Risk Matrix used in risk assesment

Sort failure procedure for operators

Instrument and room images depicting main components of ARIAIII system and room.