Versions Compared

Old Version 7

changes.mady.by.user Suat Dervish

Saved on

compared with

New Version 8

changes.mady.by.user Suat Dervish

Saved on

Key

  • This line was added.
  • This line was removed.
  • Formatting was changed.

...

  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.
    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.

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.

...

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.


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:

...

  1. 100 TRIGENE and wipe clean. Spray again with 1:

...

  1. 100 TRIGENE and allow a contact time of at least 10 minutes.

...

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.

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 1:100 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.

Notes

Dectectors

  • 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@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, 1:50 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.

Associated Documents

This Standard Operating Procedure should be read in conjunction with:

...