Method 1: Logging in Specimens and Record Keeping
Purpose
To keep a written and computerized record of all cell lines, the dates when cell lines were received and frozen, freezer locations, and any other important information such as dates of birth, sex, etc.


Procedure

1. Refer to the cell line growth record sheet. When a cell line arrives or is established from whole blood, information such as the cell line number, family position in the pedigree, sex, date of birth, date arrived, etc., is recorded on a cell line growth record sheet in a binder that corresponds to the study group to which the cell line belongs. Other information is recorded, such as the dates the cell pellet is frozen for DNA extraction and for permanent storage. The freezer locations (in stainless steel racks) of the cell line aliquots are recorded to facilitate locating the cell line at a later date.
   
2. To locate a cell line frozen for DNA extraction, first look for the location of the cell line on the master list of all the cell lines in the study group. This master list is located in the front of the study group binder to which the cell line belongs. After the rack location is identified, locate the position of the cell line in the rack. A separate binder labeled “–80 Revco” has forms representing all the racks in the freezer; refer to the –80 Revco sheet. When a cell line is removed, it is erased from the sheet and crossed off the master list in the study group binder.
   
3. A binder to locate frozen cell lines for permanent storage is labeled “–135 Cryostar”. The –135 freezer has the capacity to hold 20 racks with 10 boxes in each rack. Each box has the capacity for 81 cryotubes, with one empty space used for rack orientation. In the –135 binder, 20 dividers separate the 20 racks, and between each divider are 10 sheets (each labeled “–135 Cryostar sheet”), corresponding to the 10 boxes in that rack. The –135 Cryostar sheet is used to record the information of what cell lines are in each box. On these sheets, data such as kindred number, cell line number, and the date the cell line is frozen are recorded. When a vial is removed, it is erased off the sheet and off the master list in the front of the study group binder.
   

Method 2: Lymphocyte Transformation
Principle
Lymphocytes are transformed to establish cell lines. Mononuclear cells (lymphocytes) from anticoagulated venous blood are isolated by layering onto the histopaque. During centrifugation, erythrocytes and granulocytes are aggregated by ficoll and rapidly settle to the bottom of the tube; lymphocytes and other mononuclear cells remain at the plasma-histopaque interface. Erythrocyte contamination is neglible. Most extraneous platelets are removed by low-speed
centrifugation during the washing steps.

Special Reagents

• Cyclosporin A (CSA, from the Sandoz Research Institute; East Hanover, New Jersey 07936)
  
• Request CSA several months in advance in order to receive it when needed.
  
• Send a statement of investigator form to cover the release. It is an experimental drug and is used only in research work and not intended for human use.
  

Time Required
2–2.5 hours to prepare 2 transformations. Cell lines will require 3–4 weeks in a T-25 cm2 flask before passaging to a T-75 cm². After passaging to the larger flask, each cell line requires several more weeks to reach a cell density of 1X 108 cells/100 mL.

Procedure

1. Collect 27 mL of anticoagulated blood in 3 yellow top tubes (citrate), 9 mL each. The blood should be set up in culture as soon as possible for best results. Blood should be kept at room temperature prior to use in this procedure.
   
2. Wipe the exterior of the tubes of blood with EtOH, divide evenly, and transfer the blood into 250-mL tubes. Bring the volume of each tube up to 40 mL with wash media.
   
3. Place 10 mL of histopaque –1077 into 2 other 50-mL tubes. Overlay the blood and wash media mixture onto the 10 mL of histopaque. Do this very slowly, making sure not to mix the 2 layers.
   
4. Centrifuge tubes for 30 minutes at 1500 rpm at room temperature (no break), in the TJ-6 centrifuge. Aspirate the top layer down to within ¼" of the white blood cell layer.
   
5. Collect the WBC layer using a 10-mL pipette, moving the pipette in a circular motion around the inside of the tube just below the surface of the WBC layer. Transfer the WBC layer to another 50-mL tube.
   
6. Bring the volume of each tube up to 50 mL with wash media, gently invert tubes to mix.
   
7. Centrifuge the tubes for 20 minutes at 1200 rpm at room temperature (no break) in the TJ-6 centrifuge. Aspirate supernatant.
   
8. Add 12 mL wash media, resuspend the cell pellette, and transfer to a 15-mL centrifuge tube.
   
9. Centrifuge 8 minutes at 1000 rpm at room temperature (no break) in the TJ-6 centrifuge. Aspirate supernatant.
   
10. Cell counts can be done to determine the appropriate volume of media to be added to the cells. Cells should be set up in culture using a minimum of 2.6 × 10⁶ cells/mL and not more than 7 mL per 25 cm₂ flask. The average WBC count of whole blood ranges from 1 × 10⁶ cells/mL to 3 × 10⁶ cells/mL. An ideal primary culture should contain between 5 and 7 mL of cells/25 cm² flask. Cells are resuspended in 10 mL of RPMI (without serum) for counting. If cell counts cannot be done, set up cultures in 5–7 mL of fresh RPMI, 20% FBS, and 2 mg/mL cyclosporin A.
    
11. Inoculate cells with an equal volume of virus. Incubate cells at 37°C with 5% CO₂ and loose caps on flasks.
    
12. Feed the culture after 24–48 hours if the media has turned yellow. This is done by removing ½ of the media and replacing it with 1 mg/mL cyclosporin A media. Discontinue cyclosporin A after 3–4 weeks. Do not overfeed cells.
    Do not increase the volume of media for at least 2 weeks. If cells do not seem to be growing, reduce the volume of media and feed only once a week. After 2 weeks, when cells are very clumpy and the media is changing color from orange to yellow within 3 days incubation, increase the volume of media by 10–20 mL. Cultures are always fed by removing ½ of the old media and replacing it with a slightly increased volume of fresh media. Cultures can be split when they reach 25–30 mL of media in a 25-cm² flask.
    

Solutions

• Growth media: (600 mL)
  Add 90.0 mL FBS, 6.0 mL 200 mM (100X) L-glutamine, 0.6 mL 50 mg/mL
  gentamicin reagent to 500 mL of sterile RPMI 1640 with 2 mM L-glutamine.
  
  Filter-sterilize through a 0.22-mm filter and store up to 2 weeks at 4°C.
  
  
• Wash media: (1 liter)
  1 liter of sterile RPMI-1640 with 2 mM L-glutamine, add: 10.0 mL 2.5M
  (100X) HEPES buffer 1.2 mL 50 mg/mL gentamicin reagent.
  
  Filter-sterilize through a 0.22-mm filter and store up to 2 weeks at 4°C.
  
  
• 2X Cyclosporin A media: (1 µg/mL)
  Add 2 mL of 100X Cyclosporin A to 100 mL of growth media.
  
• 100X Cyclosporin A: (100 mL)
  Dissolve 1 mg CSA in 0.1 mL ethanol, add 0.02 mL Tween 80, and mix
  well. While continually stirring, add 1 mL RPMI, drop by drop. Quantitate
  to a final volume of 100 mL with RPMI. Filter sterilize and store at 4°C
  for up to 4 months.

Method 3: Preparation of Lymphoblastoid Cell Lines for Long-term Storage
Purpose
To store cell lines in a form that will ensure recovery with high viability. A culture in logarithmic phase of growth with a total volume of 80–100 mL/T- 75 flask should yield enough cells to freeze 10 ampules (1.0 mL/ampule). Cells should have a count of 4 X 106 cells/ampule to 9 X 106 cells/ampule. Too high or too low a cell count lowers recovery viability. Cell are frozen in RPMI-1640 with 15% Fetal Bovine Serum +10% DMSO. Cultures are frozen slowly using a Model 700 Controller freezing chamber. This precision electronic device automatically controls the injection of liquid nitrogen into the freezing chamber to provide a 1°C/minute freezing rate from +4°C to –45°C (with automatic heat of fusion compensation), then a 10°C per minute freezing rate to –90°C. Frozen ampules should be stored in liquid nitrogen for long-term storage or in a –135°C Cryopreservation System.

Cryotubes should be labeled with cell line number and date prior to the beginning this procedure.

Time Required
2.5–3.0 hours to freeze 10 aliquots from each of 6 cell lines. Only 6 cell lines or 60 cryotubes should be frozen at one time. It is essential to keep the time the cells are exposed to the DMSO at a minimum. The freezing chamber can hold up to 120 tubes so 2 people can freeze samples at the same time to save liquid nitrogen.

Procedure

1. Aspirate media from the T-75 flask down to the 50-mL mark.
   
2. Resuspend cells by shaking gently and transfer 40 mL of the cell suspension to a 50-mL centrifuge tube.
   
3. Add 10 mL of fresh media to the culture flask and reincubate at 37°C. Keep the culture flask growing until a test thaw is done on one cryotube (done to determine if the cells were successfully frozen. Refer to reactivating cell line for DNA growth and extraction procedure. The cell line will begin growing within days if the freezing conditions were correct). Greater than 99% of cell lines are successfully frozen using this procedure.
   
4. Remove 200 µL of the cell suspension from centrifuge tube for a cell count. (Refer to cell counting procedure.) Use the cell count to adjust the cell concentration to between 4 × 10⁶ and 9 × 10⁶ cells/ampule. Too high or too low a cell concentration decreases the viability of the cell line when the cryotube is thawed for growth.
   
5. Centrifuge the 50-mL tube for 10 minutes at 1200 rpm, no break, room temperature, in the TJ-6 centrifuge.
   
6. Aspirate supernatant down to ¼ inch above the cell pellet.
   
7. Place a control sample (freezing media in a 1.0-mL cryotube) into the freezing chamber in a central location, with the thermocouple probe placed equidistant from side to bottom. It will take approximately 6 minutes for the sample temperature to reach start temperature of 4°C on the chart drive.
   
8. Resuspend cell pellet with 10 mL of freezing media. Pipette 1.0 mL into each of 10 cryotubes on ice. DMSO is toxic to cells, therefore, begin freezing immediately after transferring the cells to cryotubes.
9. Load the cryotubes into the chamber when the sample temperature is +4°C on the chart drive paper.
   
10. Again allow the chamber and cells to cool to the start temperature of +4°C.
    
11. Place the selector switch to the freeze ampule position. The controller will automatically cycle through the freezing program until the end temperature is reached. This takes approximately 55 minutes.
    
12. Remove samples after the recorder has reached –90°C and transfer to a permanent storage container. Samples should be moved quickly to prevent thawing or warming and sample deterioration.
    Warning: Wear cryoprotective gloves when working with the freezing chamber and other permanent storage containers. Also, protective eyeglasses are necessary in case of the explosion of a cryotube.

Solutions

• Freezing media: (1 liter)
  Prepare a 1-liter volume and divide into 25–50 mL, centrifuge tubes
  containing 40 mL each. Store the tubes at –80°C for up to 1 year. 700 mL
  RPMI-1640 with 2 mM L-Glutamine
  
• 200 mL fetal bovine serum (FBS)
  
• 100 mL dimethyl sulfoxide (DMSO, sigma)
  
• 1000 mL total volume
  
  Filter-sterilize media and FBS with a 0.22-mm cellulose acetate filter. Do not filter DMSO; it will dissolve the cellulose acetate membrane.
  

Method 4: Reactivating Cell Lines and Cell Growth for DNA Preparation
Purpose
Cell lines are reactivated and grown to a count of 1 × 10⁸ cells. The cells are pelleted and stored frozen at –80°C prior to DNA extraction.

Time Required
15–20 minutes to begin growing 2–4 cryovials.

Procedure

1. Frozen cells should be thawed quickly. Remove the cryovial from its longterm storage container in the –135°C Cryostar, and place immediately in a 37°C water bath for 2 minutes.
   
2. Remove the cells from the vial and place in 10 mL wash media. This is necessary to remove traces of dimethyl sulfoxide from the cells.
   
3. Centrifuge cells for 10 minutes at 1200 rpm (no break) at room temperature using the TJ-6 centrifuge.
   
4. Remove the supernatant above the cell pellet.
   
5. Resuspend the cell pellet in 7–10 mL of 1X Cyclosporin A media.
   
6. Aspirate half of culture media within 3–4 days. Add growth media and slightly increase volume by 5 mL. Increase the volume of media by 5–10 mL 2 times a week by aspirating off half of media from culture flask (do not suction off cells from bottom of flask) and replacing it with fresh growth media. Cells can be harvested for extraction when a T-75 cm2 flask reaches a volume of 100 mL of media and there is a monolayer of cells on the bottom of the flask.
   

Solutions

• Wash media: (1 liter)
  Add 10.0 mL 2.5 M (100X) HEPES buffer and 1.2 mL 50 mg/mL gentamicin
  reagent to 1 liter of sterile RPMI 1640 with 2 mM L-glutamine.
  Filter-sterilize through a 0.22-mm cellulose acetate filter and store up to 2 weeks at 4°C.
  
• Growth media: (1 liter)
  Add 1 liter of sterile RPMI 1640 to 2 mM L-glutamine.
  165.0 mL fetal bovine serum, heat inactivated at 50–60°C for one and half hour 12.0 mL 200 mM (100 X) L-glutamine
  1.2 mL 50 mg/mL gentamicin reagent
  Filter-sterilize through a 0.22 µm cellulose acetate filter and store up to 2 weeks at 4°C.
  
• 1X Cyclosporin media: (100 mL)
  Add 1.0 mL 100X cyclosporin A to 100 mL of growth media.
  
• 100X Cyclosporin A: (100 mL)
  Dissolve 1 mg CSA in 0.1 mL ethanol in a sterile 15-mL centrifuge tube
  with a small magnetic stirrer. Add 0.02 mL (= 20 µL ) of Tween 80 and
  mix well. While continually stirring, add 1 mL RPMI drop by drop. Bring
  to a final volume of 100 mL with RPMI.
  Filter-sterilize with a 0.22-µm filter. Store at 4°C for up to 4 months.

Method 5: Preparation of a Lymphocyte Cell Pellet for Storage
Purpose
Following propagation to 1 X 108 cells, lymphoblastoid cells are conveniently stored at –80°C to preserve the high-molecular-weight DNA in the cells until the DNA is purified. This procedure describes the steps required to harvest and
freeze the cells for long-term storage.

Time Required
2–3 hours to prepare 12–15 cultures for storage.

Procedure

1. Aspirate the growth media from the lymphoblastoid cell culture to the 40-mL mark on the T-75 cm² flask.
   
2. Resuspend the cells in the flask by shaking gently. Remove 200 µL of the cell suspension and determine the cell count. Transfer the cell suspension either by decanting or pipetting to a 50-mL conical centrifuge tube.
   
3. Centrifuge the tubes containing cells for 10 minutes, 1200 rpm, at room temperature using the T-J6 centrifuge. Do not apply the break at the end of the centrifuge run.
   
4. Aspirate the supernatant above the cell pellet. Resuspend the cells with 10 mL of PBS
   
5. Label a 15-mL tube with the date, kindred#, cell line#, and cell count. Transfer the cell suspension to the labeled 15-mL centrifuge tube, centrifuge again for 10 minutes, and aspirate the supernatant.
   
6. Transfer the tube to a –80°C Revco freezer and record the rack location on the cell line growth record sheet.

Method 6: Maintenance of B95-8 Cell Line and Obtaining Virus for Lymphocyte Transformation
Principle
The B95-8 cell line was initiated by exposing marmoset blood leukocytes to Epstein-Barr virus (EBV) extracted from a human leukocyte line. B95-8 is a continuous line and releases high titres of transforming EBV. Thus, it provides a source of EBV to establish continuous lymphocytic cell lines from human donors.

Safety Considerations
B95-8 must be handled with precautions, since EBV can infect primates. A biological safety cabinet must be used when passaging the culture and harvesting the virus. Use bleach to kill unused virus. All material that comes in contact with the virus must be autoclaved. In addition, the door of the room should remain closed to prevent outside contaminants from entering the room and to prevent any harmful viruses from leaving the area. Gloves should always be worn in dealing with any human or hybrid cell line because latent virus genomes can be present.

Special Reagents
The B95-8 cell line. (Available from American type culture collections CAT NO. ATCC CRL 1612).

Time Required
5 minutes twice a week to feed and split the culture to maintain the correct cell density of 1.0–2.0 X 106 cells/mL.

Procedure

1. B95-8 should be grown in growth media (RPMI-1640 + 16% fetal bovine serum). The culture should be passaged twice a week: on Mondays and Thursdays, or on Tuesdays and Fridays. Passaging (subculturing) cells denotes the transplantation of cells from one culture vessel to another. Aspirate half of the old media and replace it with an equal volume of new media.
   
2. To maintain a culture at a density of around 1 X 106 cells/mL it is necessary to split it 1:4 once a week. For example, to a culture with a cell density greater than 1.5 X 106 cells/mL, one fourth is diluted with 3 parts growth media (10 mL cells +30 mL media). Save the old flask as a backup in case the new culture becomes contaminated. When the subculture is passaged the next time, dispose of the old flask.
   
3. Media containing fresh virus can be prepared at the same time the culture is passaged: Using a 10-mL or 25-mL disposable pipette, remove and transfer the media (above the cells) to a 50-mL centrifuge tube. Always be careful not to pull up any cells at bottom of the culture flask. Reserve 25 mL of media in the flask and add a equal amount of new growth media to maintain the culture.
   
4. Centrifuge the tube with the media-containing-virus for 10 minutes at 1200 rpm (no break) at room temperature, using the TJ-6 centrifuge. Centrifuging the media will pellet any marmoset cells to the bottom of the centrifuge tube.
   
5. With a 10-mL pipette, transfer all but the bottom 10 mL of virus in the centrifuge tube to a 150-mL 0.22-mm cellulose acetate filter. Filter and store the virus at 4°C for up to 7 days.
   

Solutions

• Growth media:
  Add 165.0 mL fetal bovine serum, 1.2 mL gentamicin reagent, 12.0 mL L-glutamine to 1 liter of sterile RPMI-1640.
  Filter-sterilize and store at 4°C, for up to 2 weeks.
  

Method 7: Cell Counts Using a Hemocytometer
Purpose
The purpose of this procedure is to determine the cell density of the culture. Cell cultures always have some dead cells; the viable and nonviable cells can be distinguished with the use of trypan blue dye and a hemocytometer. Living cells will not take up the dye, while dead cells do.

Time Required
5 minutes for 2 two-cell counts

Procedure

1. Transfer 200 mL of the cell suspension into a 15-mL centrifuge tube.
   
2. Add 300 mL of PBS and 500 mL of trypan blue solution to the cell suspension (creating a dilution factor of 5) in the centrifuge tube. Mix thoroughly and allow to stand 5 to 15 minutes.
   
   If cells are exposed to trypan blue for extended periods of time, viable cells may begin to take up dye as well as nonviable cells; thus, try to do cell counts within 1 hour after dye solution is added.
   
   
3. With a cover slip in place, use a Pasteur pipette and transfer a small amount of the trypan blue-cell suspension to a chamber on the hemocytometer. This is done by carefully touching the edge of the cover slip with the pipette tip and allowing the chamber to fill by capillary action. Do not overfill or underfill the chambers.
   
4. Count all the cells (nonviable cells stain blue, viable cells will remain opaque) in the 1-mm center square and the 4 corner squares. Keep a separate count of viable and nonviable cells. If more than 25% of cells are nonviable, the culture is not being maintained on the appropriate amount of media; reincubate culture and adjust the volume of media according to the confluency of the cells and the appearance of the media. A culture growing well will have many clumps of cells and will turn the media from orange to yellow within several days (increase the amount of media). A
   culture not growing well will have few clumps of cells and the media will not change to yellow (it may even turn pink); if so, decrease the volume of media). Cells may be frozen if greater than 75% of the cells are viable. Note: If greater than 10% of the cells appear clustered, repeat entire procedure, making sure the cells are dispersd by vigorous pipetting in the original cell suspension as well as the trypan blue suspension. If less than 20 or more than 100 cells are observed in the 25 squares, repeat the procedure adjusting to an appropriate dilution factor. Repeat the count using the other chamber of the hemocytometer.
   
5. Each square of the hemocytometer (with cover slip in place) represents a total volume of 0.1 mm³ or 10^–4 cm³. Since 1 cm³ is equivalent to 1 mL, the subsequent cell concentration per mL (and the total number of cells) will be determined using the following calculations.
   Cells per mL = the average count per square × the dilution factor × 104 (count 10 squares)
   Example: If the average count per square is 45 cells × 5 × 104 = 2250000
   or 2.25 × 10⁶ cells/mL.
   Total cell number = cells per mL × the original volume of fluid from which
   cell sample was removed.
   Example: 2.25 × 10⁶ (cell per mL) × 10 mL (original volume) = 2.25 × 10⁷
   total cells.

Method 8: Removal of Yeast Contamination from Lymphoblast Cultures
Purpose
This method is advantageous for saving the occasional cultures that become contaminated. Yeast-contaminated cultures will appear cloudy when slightly shaken and lymphocytes will not cluster together as much as normal. If cultures are suspect, a drop of culture can be streaked on a YPD media plate to check for growth of yeast colonies, or a 5-mL sample can be taken to Barnes Diagnostic Center for identification of yeast strain.

Procedure

1. Pipette 5 mL histopaque into a 15-mL centrifuge tube.
   
2. Carefully layer on top of the histopaque 10 mL of contaminated culture (or concentrated cells/yeast resuspended in growth media).
   
3. Centrifuge tube for 25 minutes at 2500 rpm (no break) at room temperature, using the TJ-6 centrifuge.
   
4. The yeast cells will pellet to the bottom of the histopaque gradient and the lymphoblast cells will be located on top of histopaque gradient. Remove the lymphoblast cells with a 10-mL disposable pipette, and transfer to a 15-mL centrifuge tube.
   
5. Wash cells by adding 10 mL of wash media to cells. Centrifuge 10 minutes at 1200 rpm, no break, at room temperature. Aspirate off the wash media and resuspend in RPMI-growth media containing 1X antimycotic/antibiotic.
   This will remove the rest of the yeast cells.
   
6. Transfer the cells to a 25 cm² tissue culture flask and feed the culture twice a week with 1X antimycotic/antibiotic media until all traces of contamination are gone. This will depend on the severity of the contamination (usually for cultures moderately contaminated, 2 weeks or 4 feedings will suffice). After contamination is no longer visible, feed the cultures with growth media containing only antibiotic, and not the antimycotic.
   

Solutions

• Wash media: (1 liter)
  Add 1 liter of sterile RPMI 1640 to 2 mM L-glutamine
  10.0 mL 2.5 M (100X) HEPES buffer, 1.2 mL 50 mg/mL gentamicin reagent. Filter-sterilize through a 0.22-mm cellulose acetate filter and store up to 2 weeks at 4°C.
  
• Growth media: (1 liter)
  Add 1 liter of sterile RPMI 1640 to 2 mM L-glutamine
  165.0 mL fetal bovine serum, heat inactivated at 50°C–60°C for one and half hour. 12.0 mL 200 mM (100 X) L-glutamine, 1.2 mL 50 mg/mL gentamicin reagent as added. Filter sterilize through a 0.22-µm cellulose acetate filter and store up to 2 weeks at 4°C.
  
• 1X Cyclosporin media: (100 mL)
  Add 1.0 mL 100X cyclosporin A to 100 mL of growth media.
  
• 100X Cyclosporin A: (100 mL)
  Dissolve 1 mg CSA in 0.1 mL ethanol in a sterile 15 mL centrifuge tube with a small magnetic stirrer. Add 0.02 mL (or 20 µL) of Tween 80 and mix well. While continually stirring, add 1 mL RPMI drop by drop. Quantitate to a final volume of 100 mL with RPMI. Filter-sterilize with a 0.22-µm filter. Store at 4°C for up to 4 months.
  
• Antimycotic/antibiotic media:
  Add 1 liter of sterile RPMI 1640 to 2 mM L-glutamine
  165.0 mL fetal bovine serum, heat inactivated
  12.0 mL 200 mM (100X) L-glutamine
  12.0 mL antimycotic/antibiotic (100X), liquid,
  Filter-sterilize through a 0.22-µm cellulose acetate filter and store up to 2 weeks at 4°C.
  

Method 9: Maintaining Lymphoblastoid Cell Lines
Purpose
To grow lymphoblastoid cells for permanent storage and DNA extraction. Safety Considerations All cultured animal and human cells have the potential for carrying viruses, latent viral genomes, and other infectious agents. Cell cultures should be handled very carefully by trained persons under laboratory conditions that afford adequate biohazard containment. A biological safety cabinet must be used when passaging cell lines. Use bleach in a suctioning apparatus to kill unused virus. All material used in passaging the cell lines must be autoclaved. Gloves are always worn to protect hands from contamination. A laboratory coat should be worn to protect clothes from contamination. Doors of the tissue culture room should remain closed to decrease the amount of airborn contaminants entering the incubators and the room. Equipment (incubators, centrifuges, microscopes, tabletops, etc.) should be cleaned routinely to help maintain a sterile work environment.

Time Required
3–4 weeks to grow a cell line from a frozen stock, or to grow an established cell line arriving in a T-25 cm² flask to 1 × 100 million cells.
Allow 6–8 weeks establish and grow a lymphoblastoid cell line from whole blood to 1 × 10⁸ cells.

Procedure
Maintaining lymphoblastoid cultures is fairly simple if 2 important characteristics are taken into consideration: (i) the cell cycle (primary culture and established cell line) and (ii) the cell concentration.

Cell Cycle
Every lymphoblastoid culture is unique and should be treated accordingly. For example, some cultures will grow very rapidly, while others may require twice the amount of time. Cultures that require the media to be changed every other day are rapidly dividing and will form many clumps. Cell lines which grow slowly will change the color of the media every 3–4 days, and may require the use of cyclosporin A and less media with each feeding.

Lymphoblast cultures grow in clumps and do best if periodically shaken up to break up the clumps. The cells will usually settle to the bottom of the flask, but do not attach unless the culture is in the primary stage of transformation or is lacking in nutrients. Cultures growing well will turn the media acidic within 12–24 hours after being fed. The color is a good indication of cell growth and concentration (yellow:growing well; orange or pink:not growing well).

Lymphoblasts can be grown in T-25 cm² flasks or T-75 cm² flasks. Occasionally it is necessary to use a 24 or 96 well plate if a culture is not growing well. Primary cultures are set up in a 25-cm² flask and maintained until a volume of 15–20 mL is reached. The culture is then transferred to a

Cell Concentration
The cell concentration of the suspension is important. A cell count above a certain number means decreased viability (the dead cells stain blue with Trypan blue). When the cell count is too low, cultures will show little growth. The absolute lowest cell concentration for any cell line should be 1.5–2.0 X 100 thousand viable cells/mL. Cultures can be split when the cell count is 2.0 X million viable cells/mL.

Cultures are grown upright in T-flasks. They are maintained with RPMI- 1640 (supplemented with 1% of a 200 mM L-glutamine solution) plus 15% fetal bovine serum (heat inactivated) plus an antibiotic, such as gentamicin reagent or penicillin/streptomicin. Incubation conditions are 37°C and 5% CO₂. Cultures are fed every 3 to 4 days. If a cell line is not fed frequently enough, the majority of the cells will not be in the logarithmic phase of growth; therefore, the optimum growth of the cell line is never reached. Cultures are fed by removing half of the media from the flask and replacing it with a slightly increased volume of new media. If a culture is not growing well, half of the media is removed, and the volume of added media is decreased slightly.


Method 10: Lymphoblastoid Cell Lines from Frozen Whole Blood
Purpose
Blood samples can be stored frozen as a backup in case an LCL is needed at a later date.


Time Required
15 minutes to freeze 1-4 cryotubes placing them directly into the –135°C freezer; or 1 hour to freeze the tubes using the Cryomed freezing chamber. Cells have been shown to be more viable if temperature is lowered gradually with the freezing chamber.


Procedure
Freezing cells:

1. Pipette 1.0 mL of whole blood into 2 cryotubes (1.25 mL).
   
2. Add to each cryotube 100 µL dimethyl sulfoxide (DMSO), or 10% of the volume of blood.
   
3. Immediately begin freezing the whole blood in the cryomed freezing chamber until the chart drive printer reads –90°C.
   
4. Quickly transfer the frozen sample to long-term storage in a –135°C freezer or a liquid nitrogen storage container. These whole blood samples have been shown to be viable for as long as 5 months by G. Chenevix-Trench, et al. Thawing cells for transformation:
   
5. When an LCL is needed, the cells are thawed rapidly in a 37°C water bath: Place the cryotubes in a bubble rack. Shake the rack to help thaw the cells, usually for 1–2 minutes.
   
6. With a 1-mL disposable pipette, transfer the sample to a 15-mL conical centrifuge tube filled with 10 mL of wash media.
   
7. Centrifuge the cells for 10 minutes at 1200 rpm, no break, at room temperature.
   
8. Aspirate the wash media to just above the cell pellet. Wash the pellet again with 10 mL wash media and centrifuge as in step 7. Repeat the wash a total of 4 times or until the red cell contamination is minimal. If the red cell contamination is not eliminated, several days in culture will decrease the amount of red cells substantially.
   
9. Aspirate the wash media and resuspend the cell pellet in 300 mL filtered supernatant from a B95-8 marmoset culture containing Epstein-Barr virus. Transfer the cell suspension to a T-25 cm² flask and incubate for 2 hours at 37°C with 5% CO₂. If there is a very small volume of cells, leave the cells in the 15-mL centrifuge tube for the incubation.
   
10. After incubation, add 800 mL RPMI-1640 containing 20% fetal bovine serum and 2X Cyclosporin A.
    
11. Using a 5-mL disposable pipette, plate out cells in serial dilution in a 96-well microtiter plate: transfer half of the cells in the first well into a second well. Add enough media to fill the second well. Then take half of this cell/media mixture and transfer to a third well. Fill up the third well with media. Incubate at 37°C with 5% CO₂.
    
12. Feed the cells twice-weekly by removing half of the old media and replacing with fresh media until transformed colonies are apparent (usually 2–3 weeks). The new media should contain 1X CSA.
    
13. Subculture cells to a 24-well plate before transferring to culture flasks. Maintain the subcultures on growth media (no CSA).
    

Solutions

• Growth media: (600 mL)
  To 500 mL of sterile RPMI 1640 with 2 mM L-glutamine, add: 90.0 mL FBS, 6.0 mL 200 mM (100X) L-glutamine, 0.6 mL 50 mg/mL gentamicin reagent.
  
  Filter-sterilize through a 0.22-mm filter and store up to 2 weeks at 4°C.
  
  
• Wash media: (1 liter)
  To 1 liter of sterile RPMI 1640 with 2 mM L-glutamine, add 10.0 mL 2.5 M (100X) HEPES buffer, 1.2 mL 50 mg/mL gentamicin reagent.
  
  Filter-sterilize through a 0.22-mm filter and store up to 2 weeks at 4°C.
  
  
• 2X Cyclosporin A media: (1 µg/mL)
  To 100 mL of growth media, add 2 mL of 100X Cyclosporin A.
  
• 100X Cyclosporin A: (100 mL)
  Dissolve 1 mg CSA in 0.1 mL ethanol, add 0.02 mL Tween 80, and mix well. While continually stirring, add 1 mL RPMI, drop by drop. Quantitate to a final volume of 100 mL with RPMI. Filter sterilize and store at 4°C for up to 4 months.
  

Method 11. Cell Culture Media and Solutions

Antimycotic/Antibiotic Media
To 1 liter of sterile RPMI 1640 with 2 mM L-glutamine, add:
- 165.0 mL fetal bovine serum, heat-inactivated
- 12.0 mL 200 mM (100X) L-glutamine
- 12.0 mL antimycotic/antibiotic (100X), liquid, Gibco, Cat. No. 600-5240AG
- Filter-sterilize through a 0.22-µm cellulose acetate filter and store up to 2 weeks at 4°C.

1X Cyclosporin Media: (100 mL)
To 100 mL of growth media, add 1.0 mL 100X Cyclosporin A

2X Cyclosporin A Media: (1 µg/mL)
To 100 mL of growth media add 2 mL of 100X Cyclosporin A.

100X Cyclosporin A: (100 mL)
Dissolve 1 mg CSA in 0.1 mL ethanol in a sterile 15-mL centrifuge tube with a small magnetic stirrer. Add 0.02 mL (= 20 mL) of Tween 80 and mix well. While continually stirring, add 1 mL RPMI drop by drop. Bring to a final volume of 100 mL with RPMI. Filter-sterilize with a 0.22-mm filter. Store at 4°C for up to 4 months.


Freezing Media: (1 liter)
Prepare a 1-liter volume and divide into 25–50 mL centrifuge tubes containing
40 mL each.

Store the tubes at –80°C for up to 1 year.
700 mL RPMI-1640 with 2 mM L-Glutamine
200 mL fetal bovine serum (FBS)
100 mL dimethyl sulfoxide (DMSO, Sigma)
1000 mL total volume

Filter-sterilize media and FBS with a 0.22-mm cellulose acetate filter. Do not filter DMSO, it will dissolve the cellulose acetate membrane.

Growth Media: (600 mL)

To 500 mL of sterile RPMI 1640 with 2 mM L-glutamine, add:
90.0 mL FBS
6.0 mL 200 mM (100X) L-glutamine
0.6 mL 50 mg/mL gentamicin reagent

Filter-sterilize through a 0.22-mm filter and store up to 2 weeks at 4°C.

Growth Media: (1 liter)

To 1 liter of sterile RPMI 1640 with 2 mM L-glutamine, add:
165.0 mL fetal bovine serum, heat-inactivated at 50°C–60°C for one and half hour.
12.0 mL 200 mM (100 X) L-glutamine
1.2 mL 50 mg/mL gentamicin reagent

Filter-sterilize through a 0.22-mm cellulose acetate filter and store up to 2 weeks at 4°C.

Wash Media: (1 liter)

To 1 liter of sterile RPMI 1640 with 2 mM L-glutamine, add:
10.0 mL 2.5 M (100X) Hepes buffer
1.2 mL 50 mg/mL gentamicin reagent

Filter-sterilize through a 0.22-mm cellulose acetate filter and store up to 2 weeks at 4°C.