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Protocol

IsoCode Single-Cell Adaptive Immune: TCR-T Protocol

IsoPlexis Applications Team, IsoPlexis

Aug 05, 2021

Abstract

A. Overview of Protocol

Day 1: Cryopreserved TCR-T cells are thawed and cultured overnight in the presence of IL-2. Cryopreserved target cell lines are thawed, cultured and passaged several days prior to beginning the experiment. 

Day 2: Enrichment, Staining, and Antigen Stimulation of CD4+ and/or CD8+ T cells for 20 hours. 

Day 3: Loading of TCR-T cells onto IsoCode chip. 


Introduction

NOTE: This protocol outlines the standard method for thawing and culturing of human TCR-T cells only and may not be valid for other species or cell types. 

NOTE: Using stains and protocols other than the included kit surface stains and protocols might result in failed runs. Stains and staining procedures not approved by IsoPlexis will require validation prior to use. Please consider IsoPlexis’ IsoPACE™ program to assist in custom marker and protocol validation.

Safety Warnings: Read MSDS documents of all materials prior to use. - Laboratory workers should wear standard PPE, including disposable gloves, protective eyewear, and laboratory coats. 


Reagents and Equipment

Required Consumables and Equipment


Procedure

B. Before Getting Started

1. Important Precautions

Read MSDS documents of all materials prior to use. 

Working with Biohazardous Reagents: Please refer to your institute’s guidelines and obtain proper training to handle potentially biohazardous samples. It is also strongly recommended that any lab personnel handling human samples should be vaccinated against HBV if the individual does not have sufficient HBV antibody titer. 

Additional precautions need to be taken when working with samples that potentially contain an EID agent: 

  • Laboratory workers should wear standard PPE, including disposable gloves, protective eyewear, and laboratory coats.
  • Any procedure or process that cannot be conducted in the designated EID BSC should be performed while wearing gloves, gown, goggles and a fit tested N-95 mask.
  • Work surfaces should be decontaminated on completion of work with appropriate disinfectants. This includes any surface that potentially came in contact with the specimen (centrifuge, microscope, etc.).
  • All liquid waste produced in the processes must be treated to a final concentration of 10% bleach prior to disposal.

2. Reagents to Be Prepared Before Starting

C. Protocol 

Please read section B before starting protocol. 

Note: All incubation and centrifugation steps are performed at room temperature unless specified otherwise. 

Prior to Day 1: Thawing and Culturing of Target Cell Lines 

Note: If required to expand target cell lines, this should be done several days in advance of Day 1. 

Thawing and Culturing of Target Cell Lines:

 1. Aliquot 5 mL of pre-warmed complete RPMI into a 15 mL Falcon centrifuge tube. 

2. Using proper PPE, remove selected vial(s) from liquid nitrogen storage. Quickly remove to water bath (37°C) to thaw. While thawing, swirl in the water until a single ice crystal remains in the vial. Be sure to prevent (to the best of your ability) any of the water from the water bath to get underneath the cap and into the sample. 

3. When the sample is nearly thawed, immediately spray vial with 70% alcohol before bringing into the hood. It is important to allow the alcohol to evaporate before opening the vial. 

4. As soon as the vial is dry, add 1 mL of pre-warmed complete RPMI growth media to the vial. Pipette up and down to mix and transfer the sample to the prepared centrifuge tube (with the warmed media), washing the inside of the cryogenic vial with the media from the tube as well. Centrifuge at 300xg (RCF) for 10 minutes and aspirate supernatant. 

5. Depending on the number of cells cultured, resuspend the pellet to a density of 1 x 106 cells/mL in complete RPMI so that the final RPMI volume is 3-5 mL (T25) or 8-10 mL (T75) and transfer to the respective flask. Incubate at 37°C, 5% CO2. 

6. Passage cells every few days depending on the requirements for your specific cell type. 

Note: All liquid waste produced in this process must be treated to a final concentration of 10% bleach prior to disposal. 

Day 1: Culture of Target Cells and Recovery of TCR-T Cells 

Culturing of Target Cells:

1. Passage the target and control cells needed for the experiment into a T75 at a concentration of 1 x 106 cells/mL. 

Note: Passage sufficient numbers of target and control cells to perform assay at a ratio of 1:2 TCR-T cells to targets. 

2. Incubate overnight at 37°C, 5% CO2. 

Note: All liquid waste produced in this process must be treated to a final concentration of 10% bleach prior to disposal. 

Thawing and Culturing Cryopreserved TCR-T Cells:

1. Aliquot 8 mL of pre-warmed complete RPMI into a 15 mL Falcon centrifuge tube. 

2. Using proper PPE, remove selected vial(s) from liquid nitrogen storage. Quickly move to water bath (37°C) to thaw. While thawing, swirl in the water until a single ice crystal remains in the vial. Be sure to prevent (to the best of your ability) any of the water from the water bath to get underneath the cap and into the sample. 

3. When the sample is nearly thawed, immediately spray vial with 70% alcohol before bringing into the hood. It is important to allow the alcohol to evaporate before opening the vial. 

4. As soon as the vial is dry, add 1 mL of pre-warmed complete centrifuge tube (with the warmed media), washing the inside of the cryogenic vial with the media from the tube as well. Centrifuge at 300xg (RCF) for 10 minutes and aspirate supernatant. 

5. Depending on number of cells cultured, resuspend the pellet to a density of 1 x 106 cells/mL in complete RPMI supplemented with 10 ng/mL IL-2 (equivalent of 236 IU/mL) so that the final RPMI volume is 3-5 mL (T25) or 8-10 mL (T75) and transfer to the respective flask. 

6. Incubate overnight at 37°C, 5% CO2. 

Day 2: Sample Enrichment 

Summary: 

After IL-2 recovery, CD8 T-Cells are enriched by anti-CD8 microbeads. The flow through will be used as the CD4 fraction. Target cells are also pulsed with peptide(s) for 2 hrs before co-culture. 

Note: Standard practice is to enrich the smaller fraction first. Generally, this is the CD8 fraction however this may be different depending on your specific samples. Note: This protocol describes enrichment of one fraction with the depleted flow through serving as the other fraction. 

1. Harvest TCR-T cell culture(s) by gently pipetting up and down using pipettor and transfer to 15 mL falcon tube. 

2. Count cells using a hemocytometer or automated cell counter and determine percent of viable cells as described in Appendix D1. 

3. Perform dead cell depletion if the number of dead cells is >20%. Please refer to “Protocol: Dead Cell Removal Using Ficoll” found in Appendix D2 of this protocol. 

4. Centrifuge cells at 300xg (RCF) for 10 minutes. 

5. Aspirate supernatant completely. 

6. Resuspend cell pellet in 80 µL of cold (2-8°C) RoboSep (Stem Cell Technologies) for up to 107 cells, e.g., use 160 µL for 2 x 107 cells. 

7. Add 20 µL of CD8 MicroBeads (Miltenyi) for up to 107 cells, e.g., use 40 µL for 2 x 107 cells. 

8. Mix well and incubate for 15 minutes in the refrigerator (2-8°C). 

9. Wash cells by adding 1-2 mL of cold (2-8°C) RoboSep for up to 107 cells. 

10. Centrifuge at 300xg (RCF) for 10 minutes. 

11. Aspirate supernatant completely. 

12. Resuspend 108 cells or fewer in 500 µL of cold (2-8°C) RoboSep. 

13. Place LS column in the magnetic field of the MACS separator with the wings facing out. 

14. Rinse column with 3 mL of cold (2-8C) RoboSep buffer into a 15 mL Falcon tube and discard. 

15. Place new 15 mL Falcon tube underneath column. 

16. Pipette cell suspension from Sample Enrichment step 12 onto the column. 

17. Collect unlabeled cells: Let the column reservoir empty by gravity and then flush out unbound cells by adding 3 mL of cold (2-8°C) RoboSep buffer. 

18. Repeat step 17 two more times for a total of 3 washes while collecting into the same Falcon tube. This is the CD8-depleted cell fraction that contains CD4+ T cells. Do not discard this tube as it will be used as your CD4+ T cell subset in Cell Staining Step 1. 

19. Collect positively selected cells: Remove column from the magnetic separator and place on a new 15 mL Falcon tube. 

20. Pipette 5 mL cold (2-8°C) RoboSep buffer onto the column. Immediately flush out the magnetically labeled cells by firmly pushing the plunger into the column reservoir. This is your CD8+ cell fraction. 

21. OPTIONAL: To increase number of recovered cells: Remove plunger and pipette another 3-4 mL of cold (2- 8°C) RoboSep onto the column and immediately flush out any remaining magnetically labeled cells (into same collection tube) by firmly pushing the plunger into the column. 

Day 2: Cell Staining 

1. Prepare membrane stain stock.

  • Thaw tube of membrane stain diluent (DMSO) at room temperature.
  • Spin tubes of membrane stain and membrane stain diluent (DMSO) in a mini centrifuge for 10 seconds to collect the contents at the bottom of the tubes.
  • Add 20 µL of membrane stain diluent (DMSO) directly to the tube of membrane stain. Pipet up and down 15 times gently to resuspend. 

CRITICAL: Membrane stain must be prepared fresh. Discard remaining stain – do not store. 

2. Prepare stain master mix by diluting 2 µL of membrane stain into 1 mL of 1X PBS in a Lo-Bind Eppendorf tube (1:500 final dilution). With the same pipette tip, pipette up and down 10 times to ensure all membrane stain has been released. Depending on sample number and cell count, additional tubes of stain master mix may need to be prepared. CRITICAL: Failure to follow these steps will negatively impact cell counts.

  • With a P1000 set to 500 µL, gently pipette the stain master mix up and down 15 times.
  • Gently vortex the stain master mix for 5 seconds.
  • Ensure master mix is mixed well before adding stain to cells. 

3. Centrifuge enriched cell fractions at 300xg (RCF) for 10 minutes and aspirate supernatant. 

4. Add 1 mL of PBS to dilute any remaining media and mix by pipetting up and down. 

5. Centrifuge cells for 10 minutes at 300 rcf. 

6. After cells are centrifuged, check for cell pellets. 

7. Aspirate supernatant with a pipette.* TIP: Be careful not to aspirate the cell pellets.

8. For every 1 x 106 cells, add 100 µL of well mixed stain master mix to each cell suspension tube. CRITICAL: Pipet mix the cells 15 times. Be careful to not create bubbles. Gently remix master mix if it has been sitting for longer than a few minutes. 

9. Incubate for 5 minutes at 37°C in the dark. 

10. Gently pipet mix the cell suspension 15 times. CRITICAL: Be careful to not create bubbles. 

11. Incubate for an additional 5 minutes at 37°C in the dark. 

12. After incubation, add 5 times the volume of complete RPMI. CRITICAL: Pipet mix the cells 15 times. Be careful to not create bubbles. 

13. Incubate for 10 minutes at 37°C in the dark. 

14. Take an aliquot of cells to count (10 µL). See Appendix D1 for cell counting protocol. 

15. Centrifuge the stained cell fractions at 300xg (RCF) for 10 minutes.

16. Aspirate the supernatant and resuspend the T cells at 1 x 106 cells/mL in complete RPMI. Keep cells in the incubator at 37°C 5% CO2 until co-culture step. 

Note: Cell stimulation protocol is dependent upon target cell type (adherent vs. non-adherent). Proceed. To the appropriate protocol below. 

Day 2: Cell Stimulation (Adherent) 

1. Remove media from flask containing adherent cells. Discard this supernatant as viable cells should be attached to flask. 

2. Wash adherent cells with PBS. Remove PBS and discard. 

3. Detach cells with Trypsin solution following the manufacturer’s suggested instructions for specific cell types. 

4. Rinse detached cells with complete RPMI media and transfer to 15 mL Falcon tube. 

5. Count cells using a hemocytometer and determine percent of viable cells as described in Appendix D1 of this protocol. 

6. Centrifuge cells at 300xg (RCF) for 10 minutes and aspirate supernatant. 

7. Resuspend adherent target cells in complete RPMI media at a density of 1 x 106 /mL. 

8. To pulse cells, incubate target cells with desired concentration of peptide(s) in a cell culture plate for 2 hours at 37°C, 5% CO2. Use Table 7 to determine appropriate plate for cell suspension volume. Mix cells by pipetting up and down every 30 minutes. 

9. After 2 hours, lift cells by gentle scraping, if needed. 

10. Rinse wells with PBS (volume according to Table 7) and transfer to 15 mL Falcon tube. 

11. Centrifuge the pulsed and unpulsed target cells at 300xg for 10 minutes, aspirate the supernatant and resuspend in complete RPMI media at a density of 2 x 106 /mL. 

12. Seed cell suspension into a well of an appropriately sized plate as per Table 7. 

13. Culture cells at 37°C, 5% CO2 for 2 hours until cells are beginning to adhere to plate. 

14. To the cell culture plate seeded in Step 13, add CD8+ TCR-T cells or CD4+ TCR-T cells from Cell Staining Step 7 (concentration: 1 x 106 cells/mL) at a 1:1 volume ratio for an overall density ratio of 1 TCR-T cell to 2 target cells. 

15. Centrifuge the plate at 300xg for 5 minutes to bring TCR-T cells to bottom of wells. 

16. Incubate the co-culture at 37°C, 5% CO2, for 20 hours. 

Day 2: Cell Stimulation (Non-adherent) 

1. Harvest target cell culture(s) by gently pipetting up and down using pipettor and transfer to 15 mL Falcon tube. 

2. Count cells using a hemocytometer and determine percent of viable cells as described in Appendix D1 of this protocol. 

3. Perform dead cell depletion if the number of dead cells is > 20%. Please refer to “Protocol: Dead Cell Removal Using Ficoll” found in the Appendix D of this protocol. 

4. Centrifuge cells at 300xg (RCF) for 10 minutes. Aspirate supernatant. 

5. Resuspend target cells to be pulsed with complete RPMI at a density of 5-10 x 106 /mL and transfer to a 5 mL round bottom polypropylene tube. 

6. To pulse cells, incubate target cells in the round bottom tube with desired concentration of peptide(s) for 2 hours at 37°C, 5% CO2. 

7. Mix the cells by pipetting up and down every 30 minutes. 

8. After 2 hours, harvest pulsed target cell culture(s) by gently pipetting up and down using a pipettor and transfer to 15 mL Falcon tube. 

9. Rinse cells once with the addition of 5 mL of complete RPMI media. Mix well by pipetting up and down. 

10. Take an aliquot of cells to count (10 µL). Count cells using a hemocytometer and determine the percent of viable cells as described in Appendix D1 of this protocol. 

11. Centrifuge cells at 300xg (RCF) for 10 minutes. 

12. Aspirate the supernatant and resuspend peptide-pulsed and unpulsed target cells in fresh complete RPMI media at a density of 2 x 106 /mL. 

13. Mix CD8+ TCR-T cells or CD4+ TCR-T cells from Cell Staining Step 7 (concentration: 1 x 106 cells/mL) with pulsed or unpulsed target cells (concentration 2 x 106 cells/mL) in a 96 well U-bottom plate, at a 1:1 volume ratio (100 µL target cell suspension and 100 µL TCR-T cell suspension) for an overall density ratio of 1 TCRT cell to 2 target cells. Final volume in each well should be 200 µL. 

14. Incubate the co-culture at 37°C, 5% CO2 for 20 hours. 

Note: Loading cells protocol is dependent upon target cell type (adherent vs. non-adherent). Proceed to the appropriate protocol below. 

Day 3: Loading Cells (Adherent) 

1. After stimulation, collect TCR-T cells from culture by gently pipetting up and down without disturbing the adherent targets. 

CAUTION: Be careful not to lift the adherent target cells with TCR-T cells. 

2. Transfer TCR-T cells to a Lo-Bind microcentrifuge tube. 

3. Centrifuge for 10 minutes at 300xg (RCF) and collect supernatant.* 

4. Resuspend the cells in 500 µL of complete RPMI. 

5. Take 10 µL aliquot and count live cells as described in Appendix D1 in this protocol. 

6. Adjust cell concentration with complete RPMI to a cell density of 1 x 106 cells/mL and load immediately onto the IsoCode chip using 30 µL of well-suspended cell solution. Please refer to the loading instructions of the IsoLight instrument for details. 

* Note: OPTIONAL: Collect supernatants (100 µL per well) from all groups and store at -80°C for population control. 

Day 3: Loading Cells (Non-adherent) 

1. After stimulation, transfer stimulated cell culture from 96 well plate to a Lo-Bind microcentrifuge tube. 

2. Centrifuge for 10 minutes at 300xg (RCF) and collect supernatant.* 

3. Enrich TCR-T cells by removal of target cells using antibody-conjugated magnetic beads specific to target cells using Cell Depletion Protocol as described in Appendix D3. 

4. Following Cell Depletion Protocol, cells should be at a cell density of 1 x 106 . Load immediately onto the IsoCode chip using 30 µL of well-suspended cell solution. Please refer to the loading instructions of the IsoLight instrument for details. 

*Note: OPTIONAL: Collect supernatants (100 µL per well) from all groups and store at -80°C for population control. 

 


Notes and Comments

D: Appendix

1. Protocol: Cell Quality Control (CQC) 

1. Take small amount of sample (e.g., 6-10 µL) and add equal volume of Trypan blue solution. Mix gently and load onto hemocytometer. Repeat count if more than 200 cells/16 squares were counted using a 1:5 or 1:10 dilution with PBS or media using a fresh sample aliquot. 

2. Count and record viable (clear) and dead cells (blue) of all four 16-square corners. 

3. Calculate the concentration of cells as follows:

  • Concentration (cells/mL) = Average per square cell count x 104 x dilution factor

4. Calculate the number of cells as follows:

  • Number of cells = Cell concentration (cells/mL) from D.1.3 x total volume of cell suspension (mL)

5. Calculate percent viable cells:

  • % Viable cells = 100 x number of viable cells/[number of viable cells + number of dead cells]

2. Protocol: Dead Cell Removal Using Ficoll 

Note: It is recommended to start with a minimum of 3 x 106 cells. 

1. Prepare Ficoll tubes to deplete dead cells: Carefully add 6 mL of Ficoll to the bottom of the required amount of 15 mL Falcon tubes prior to harvesting stimulation cultures. 

2. Centrifuge cells at 300xg (RCF) for 10 minutes. Aspirate supernatant and resuspend the pellet in 7 mL of RPMI. 

Note: Do not use more than 1 x 107 cells of your suspension per Ficoll tube. 

3. Add the cell suspension VERY SLOWLY to the tubes containing Ficoll by placing the tip of your pipette on the wall of the tube, close to the Ficoll layer. 

CAUTION: This step must be done carefully and slowly to avoid mixing of the layers. 

4. Centrifuge tubes at 300xg for 20 minutes in a swinging bucket rotor at room temperature with acceleration and brakes off to preserve the density layers established during centrifugation. 

5. Prepare appropriate amount of Falcon tubes containing 6-10 mL of RPMI while waiting for the centrifugation to complete. 

6. Collect the viable cells by recovering the cloudy layer between Ficoll and complete RPMI media and place cells in the 15 mL Falcon tube containing media. 

7. Count cells as described in Appendix D1. 

3. Protocol: Cell Depletion

1. Remove depletion beads from the refrigerator. Resuspend depletion beads using a P1000 pipet. 

2. For every 3 x 105 cells, add 50 µL of depletion beads (e.g., antiCD235a-conjugated beads for K562 cell depletion) to each cell pellet and return remainder of beads to 4°C. 

3. Incubate for 10 minutes at room temperature, gently mix with a P100 pipet to ensure beads are kept in suspension. Resuspend every 2 minutes to keep mixture in suspension. 

4. After the incubation, add 950 µL of PBS to the Lo-Bind microcentrifuge tube. 

5. Gently pipet up and down to resuspend. 

6. Transfer mixture from Lo-Bind microcentrifuge tube to a pre-labeled 5 mL polystyrene tube (provided in Cell Line Depletion kit). 

7. Place 5 mL polystyrene tube in an EasySepTM magnet for 2 minutes. 

8. After 2 minutes, keep 5 mL polystyrene tube attached to magnet and forcefully decant T cells in a 15 mL conical tube being careful not to disrupt the 5 mL polystyrene tube. 

9. Remove 5 mL polystyrene tube from magnet. 

10. Add 1 mL of PBS to the empty Lo-Bind microcentrifuge tube from Step 10 to recover any remaining material. Transfer contents to corresponding 5 mL polystyrene tube. Be sure to rinse the walls of 5 mL polystyrene tube well and resuspend beads. 

11. Place 5 mL polystyrene tube in an EasySepTM magnet for 2 minutes. 

12. After 2 minutes, keep 5 mL polystyrene tube attached to magnet and forcefully decant T cells in a 15 mL conical tube being careful not to disrupt the 5 mL polystyrene tube. 

Note: Pool target cell depleted T cell suspensions from the same sample into one 15 mL conical tube. 

13. Centrifuge T cells collected in the 15 mL conical tube at 300xg (RCF) for 10 minutes. 

14. Remove supernatant and resuspend in 100 µL of complete RPMI. 

15. Remove 10 µL aliquot for cell counting as described in Appendix D1. 

16. Centrifuge T cells at 300xg (RCF) for 10 minutes. 

17. Remove supernatant and resuspend in an appropriate volume of complete RPMI media for a cell density of 1 x 106 cells/mL.

For troubleshooting, contact Support at 475-221-8402 & support@isoplexis.com.

 


References

IsoPlexis


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