Table of Contents
A. Obtaining Media
B. Making Supplement Stocks
- BPE (Bovine Pituitary Extract)
- EGF (Epidermal Growth Factor)
- Transferrin, Human
- Insulin
- Hydrocortisone
- Cholera Toxin
- Isoproteranol
- Oxytocin
- E2 (ß-estradiol)
- T3 (Triiodo-L-thyronine)
- Albumax
- BSA
- L-glutamine
- Penicillin/streptomycin
- Making complete media with supplements
II. Growing and Subculturing Cells in M87A (serum-containing) Medium
III. Growing and Subculturing Cells in Serum-Free Medium (MCDB 170-like)
IV. Tissue Processing Protocol
A. Tissue Processing
B. Seeding Frozen Organoids
C. Subculture of Primary Organoid Cultures
D. Selection of Cells in MCDB 170
E. Propagation of Cells in MM
(references: Garbe et al. 2009, LaBarge et al. 2103, Hammond et al. 1984, Stampfer 1985)
Basal M87A can be purchased from the UCSF Cell Culture Facility. Contact Amy Choi, media production supervisor, at amy.choi@ucsf.edu; 415-476-8686. You will need to add supplements before use. If you want to make up your own M87A please contact us for more information.
MCDB 170-like media can be purchased from Lonza (MEBM, MEGM) or ThermoFisher (HuMEC medium, Medium 171).
Maintenance of the appropriate pH is critically important. Media color should be salmon-pink. If it gets too acid or basic it will diminish the cells’ growth potential.
Fetal calf serum (we order from ThermoFisher Scientific; cat# 26140)
100 ml bottles of FBS can be thawed and kept at 4°C for ongoing use. Larger volumes can be aliquoted and refrozen for storage at -20°C/-80°C.
BPE (Bovine Pituitary Extract): we order Hammond Cell Tech cat#1078-NZ
Stock BPE is at ~14 mg/ml. Final concentration in medium is ~35 µg/ml.
Stable for 2 years
In our experience BPE is best when stored at -80°C. We’ve seen it deteriorate even when stored at -20°C. The best quality BPE will have a color (and odor) reminiscent of raw steak.
Other sources of BPE (Bovine Pituitary Extract):
JR Scientific (50008-50; 50 ml non-sterile crude preparation at up to 20 mg/ml)
Lonza (cat# cc-4009; 2 ml at 13 mg/ml)
BDBiosciences (cat# 356123; 75 mg lyophilized)
ThermoFisher Scientific (cat# 13028014; 25 mg liquid)
AthenaES (cat# 0413; 100 mg at 9-20 mg/ml liquid)
Millipore (cat# 02-103; 50 mg lyophilized)
EGF (Epidermal Growth Factor, human recombinant): Upstate Biotechnology: cat# 01-107 or ThermoFisher Scientific cat# PHG0311
Stock is 0.1 mg/ml. Final concentration in medium: 5 ng/ml.
• Add 1 ml sterile dH2O to 100 µg EGF.
• Aliquot into sterile ampoules.
Store at -20°C. Avoid repeated freeze/thaw cycles.
Apo-transferrin, Human: SIGMA cat# T2252
Stock is 10 mg/ml. Final concentration in medium is 2.5 µg/ml.
• Dissolve 100 mg of apo-transferrin into 10 mL distilled water.
• Filter sterilize through 0.2µ filter.
• Aliquot into ampoules.
Store at -20°C. Avoid repeated freeze/thaw cycles. Stable for 1 year.
Insulin: SIGMA cat# I5500
Stock is 5 mg/ml. Final concentration in medium is 7.5 µg/ml.
* If the solution is not clear after a reasonable amount of stirring, add a few more µl of 1 N HCl. (The total [HCl] should not exceed 0.005 N HCl). When the solution clears, then bring up to 100 ml with distilled water.
Avoid repeated freeze/thaw cycles. Stable for 1 year.
NOTE: It may be difficult to obtain bovine pancreatic insulin. Our testing has shown that human recombinant insulin (100 ng/ml) plus recombinant IGF-I (10 ng/ml) can be used as a replacement.
Insulin-human recombinant SIGMA cat#I2643
Stock is 1 mg/ml. Final concentration in medium is 100 ng/ml.
* If the solution is not clear after a reasonable amount of stirring, add a few more µl of 1 N HCl. (The total [HCl] should not exceed 0.005 N HCl). When the solution clears, then bring up to 50 ml with distilled water.
Avoid repeated freeze/thaw cycles. Stable for 1 year.
IGF-1, human recombinant (Long® R3-IGF-1) SIGMA cat#I1271
Stock solution is 100 µg/ml. Final concentration is medium is 10 ng/ml.
Stock solutions of peptide can be stored for at least 3 months at –20°C or –70°C.
Avoid repeated freeze-thaw cycles.
Hydrocortisone: SIGMA cat# H4001
Stock is 1 mg/ ml. Final concentration in medium: 0.3 µg/ ml.
• Dissolve 50 mg hydrocortisone in 50 ml of absolute EtOH.
Store at -20 C. Stable for 1 year.
Cholera Toxin: SIGMA cat# C8052
Stock is 10 µg/ml. Final concentration in medium is 0.5 ng/ml.
• Add 0.5 ml dH2O to 0.5 mg cholera toxin bottle to make 1 mg/ml.
• Take 0.1 ml and dilute in 10 ml sterile dH2O.
• Aliquot into 1 ml ampoules.
Store in refrigerator. Do not freeze. Stocks stored in refrigerator are stable for 3 months (Sigma says stable 1 year).
(±)Isoproterenol: SIGMA cat# I5627 (a.k.a. Isoprenaline hydrochloride)
Stock is 5X10-3 M. Final concentration in medium is 5x10-6.
DO NOT BREATHE DUST.
• Dissolve 50 mg (±)isoproterenol in 40 ml of 95% ethanol.
Store at -20°C. Make up fresh each month.
Oxytocin: Bachem cat# H2510
Stock is 1 µM. Final concentration in medium is 0.1 nM.
• Dissolve 5 mg of oxytocin peptide in 5 ml PBS + 1% BSA to give a stock of 1 mM.
• Aliquot 50 µl of stock into microfuge tubes and lyophilize/speed vacuum dry (peptides are more stable when dry than in solution).
• Store dry peptide at -20°C.
• To reconstitute oxytocin, add 500 µl PBS to dried peptide to make a 100 µM solution.
• Dilute 100µl of this into 10 ml of PBS + 0.1% BSA and filter sterilize to make 1.0 µM working stock.
• Aliquot into sterile ampoules.
Store at -20°C. Make fresh working stock every 6 months. Avoid repeated freeze-thaw cycles.
b-Estradiol: (E2) SIGMA cat# E8875
Stock is 2 x 10-5 M. Final concentration in medium: 5x10-10 M.
• Dissolve 5.44 mg in 10ml absolute EtOH to make 2 x 10-3M solution.
• Dilute 100µl of this into 10ml for a 2x 10-5 M stock.
Store at -20°C.
3,3',5-triiodo-L-thyronine: (T3) SIGMA cat# T2877
Stock is 2 x 10-4 M. Final concentration in medium: 5x10-9 M.
• Add 1.302 mg tri-iodo-thyronine to 10 ml dH2O.
• Add 1 drop 0.1 M NaOH and allow to dissolve.
• Filter sterilize and make 1 ml aliquots.
Store aliquots at -20°C. Thawed aliquots can be stored at 4°C for 1 month.
AlbuMAX I: (lipid rich bovine serum albumin): ThermoFisher Scientific cat#11020-021
Final concentration in medium is 0.1%.
Albumax I can be added as a powder followed by filtration of the final medium. If you choose, make a 20% stock in DME:F12 basal medium and filter sterilize. Make aliquots and store at -20°C. Add 2.5 ml per 500 ml of medium. Stable for 1 year as a frozen solution. We make a 10% solution in the 100x glutamine stock, which is then sterilized by filtration. Adding 5 mls of this solution gives 0.1% Albumax and 2 mM glutamine.
BSA: (bovine serum albumin): SIGMA cat# A4161
This is used to prepare the oxytocin (and IGF-1 if used). Stock is 2%. Final concentration used is 0.1%.
• Dissolve 2 g of BSA into 100 ml PBS.
• Filter sterilize through 0.2µ filter.
Store at 4°C for 6 months or aliquot and store at -20°C for 2 years.
L-glutamine: 200 mM (we order from ThermoFisher Scientific; cat# 25030)
Purchased as a 200 mM stock solution in 100 ml bottles. It is purchased and stored frozen at -20°C. Once thawed a botttle can be kept for 1 month.
Penicillin/streptomycin: (we order from ThermoFisher Scientific; cat# 15070)
Purchased as a frozen 100X stock solution in 100 ml bottles and stored at -20°C. Once thawed it is good for over 1 year. It can be aliquoted and refrozen if desired.
We do not routinely add Pen/Strep to our cultures. As a cell bank, we want to be aware of any breach of sterility that could also involve mycoplasma. However, for short-term experiments or cultures that will not be banked, adding Pen/Strep is fine.
Antibiotics for selection
Puromycin 1 µg/ml for selection; 0.5-1.0 µg/ml for maintenence
Hygromycin 10 µg/ml for selection; 5 µg/ml for maintenence
G418 100 µg/ml for selection; 50 µg/ml for maintenence
Blasticidin-S 10 µg/ml for selection; 1-5 µg/ml for maintenence
To make 500 ml of complete M87A medium with Cholera Toxin and Oxytocin:
Add the following supplements to one 500 ml bottle of M87A.
Factor Amount |
Glutamine** 5.0 ml |
Fetal bovine serum 1.25 ml |
Bovine pituitary extract 1.25 ml |
Insulin 0.75 ml |
Isoproterenol 0.5 ml |
Hydrocortisone 0.15 ml |
Apo-transferrin 0.125 ml |
Oxytocin 0.05 ml |
Cholera toxin 0.025 ml |
Epidermal growth factor 0.025 ml |
β-estradiol 0.0125 ml |
Tri-iodo-thyronine 0.0125 ml |
Albumax I ** 0.5 gm |
**Albumax I can be added as a powder followed by filtration of the final medium or can be added as 2.5 ml of a 20% stock solution. We make a 10% solution in the 100x glutamine stock, which is then sterilized by filtration. Adding 5 mls of this solution gives the desired concentration of Albumax and glutamine.
(references: Garbe et al. 2009; LaBarge et al. 2013; Stampfer, 1982; Stampfer 1985)
Dish/Flask |
Amount Media (ml) |
Number of Cells for Seeding |
T-75 |
12-15 |
2-5 x 10e5 |
100mm |
10-12 |
2-5 x 10e5 |
60mm; T-25 |
4-5 |
1.0-1.5 x10e5 |
35mm |
1.5 |
0.5 x 10e5 |
24 Well Plate |
1.0/well |
0.2 x 10e5/well |
Cells plated at these densities will increase 6-10x in number before confluence. We routinely grow cells in dishes, not flasks, because of the better gas exchange. We only use flasks if cells need to be transported.
M87A does not support clonal growth of HMEC, so seeding at densities lower than those indicated can lead to reduced cell viability. Cells should be subcultured when subconfluent to just confluent; once confluent, viability will rapidly diminish. When we re-feed cultures, we usually leave a little of the spent media.
1) Label and add medium to all dishes.
2) Put 2 scoops of liquid nitrogen or dry ice into styrofoam box containing ampoule rack and cover with lid.
3) Remove cells from freezer and immediately place in box, cover with lid.
4) Thaw one ampoule at a time in 37°C water bath, do not immerse top into water. As soon as it is thawed (do not leave in bath), wipe down with 70% ethanol in the hood.
5) Open amp and dilute into tube with appropriate volume of medium for seeding; mix well with media for a good distribution. i.e., if seeding 5x10e5 cells into 3-60's, place the 0.5 ml already in the amp into a tube with either 1 or 2.5 ml of media, mix, and distribute either 0.5 or 1 ml to each 60 mm dish containing 4 ml of medium. It's most important to mix cells well and distribute cells evenly.
6) Distribute cells evenly by gently pushing dish side-to-side up-and-down 5-10 times in each direction. Check distribution under the microscope. Place in incubator.
We do not pellet cells thawed from the freezer and we do not recommend that you do either. Cells are frozen in 1:1 DME/F-12 with 44% FCS and 6% DMSO (CPMII).
It is important to subculture the cells when they are subconfluent or just confluent, as they lose viability when kept in confluent cultures. We do most of our experiments, or harvest cells for RNA and DNA, when they are approaching subconfluence and still actively proliferating, and the day after feeding (to maintain cell-cycle consistency). If it is important to have randomly cycling cells, feed 72, 48, and 24 hrs prior to harvesting or subculture. When the cells are growing as they should, they are subcultured around every 4-7 days at 1/6 to 1/10 split ratios. If your cells are not growing this fast, something is wrong. Call!
We do not subculture all cells of one specimen at the same time. e.g., if there are 3 60s, we subculture 1 one day, and wait 48 hrs before discarding the extra dishes, or, wait 24 hrs to freeze the extra dishes. If you must split all the cells the same day, I recommend doing it in two separate, (non-cross contaminatable) batches.
NOTE ON TRYPSIN: Always label trypsin with the date thawed; do not use trypsin more than one week old. Do not forget to take trypsin out of 37°C water bath immediately when thawed. Better yet, give it the time to thaw at room temperature. Store at 4°C.
The most common error I have observed is not following the instructions for subculture. DO NOT IMPROVISE! Follow these instructions. These cells are very tightly attached to the substratum and need strong measures to be removed.
Need:
1) Look at cells under microscope. Make sure they are not contaminated and look appropriate.
2) Aspirate all media from parent dish. Wash once with STV (not something else):
3) Add just enough STV to barely cover the cells (important to minimize amount of STV), in general:
4) Incubate cells in 37°C incubator for 2-5 minutes. This time is not fixed, as cells vary. Do not leave at room temperature. The cells will not come off at room temperature.
5) After ~2’ take cells out of incubator and check under microscope to see if they have "rounded up". Tap dish lightly but sharply against microscope or desk if necessary to knock cells loose. Most (not all) cultures will still be largely attached, and you can guesstimate how long they will need to stay in the trypsin to be detached. Don't leave the STV on longer than necessary to remove most (don't need to wait for 100% if some cells remain stuck) of the cells - trypsin chews up cells. Continue incubation at 37°C, with periodic checks under the microscope, until most cells are detached.
6) When most of the cells have loosened add M87A to culture vessel, preferably using a plugged Pasteur pipette with a hand pipetter, or otherwise regular pipettes. Do not wait for all the cells to come off, particularly if only small patches remain. Repipette the medium in the flask/dish to break up clumps, and then transfer this to 15 or 50 ml tubes (use the larger volume tube for 100mm dishes, T-75, or more than 2-60mm dishes). Wash the flask/dish with about the same amount of PBS one or two more times and transfer to tube. The final volume in the tube should be about:
depending upon the cell density of the culture
7) Repipette to mix and to break up any cell clumps to facilitate counting single cells. Take a small amount in the tip of a plugged Pasteur pipette and drop onto both sides of a haemocytometer. Note the volume in the centrifuge tube.
8) Check under microscope that both chambers of haemocytometer have approximately the same number of cells (low power). The ideal # of cells to count is about 100 cells/5 squares/chamber.
9) Count cells in haemocytometer (10X power). We count 5 squares per chamber x 2 chambers. Record # of cells counted/chamber, and determine total cell count (using the noted volume in the centrifuge tube). Based on what's to be done with cells, calculate dilutions as appropriate for seeding. We generally dilute to about 10e5 cells/ml. Label dishes/flasks to be seeded and add necessary amount of medium.
10) Seed dishes and distribute cells evenly by gently pushing dish side-to-side up-and-down 5-10 times in each direction. Check distribution under the microscope. Place in incubator.
We always freeze our cells at the density of 10e6 cells/ml freeze medium. Cells are generally frozen in 0.5 or 1.0 ml amounts (5 x 10e5 or 10e6 cells), depending on number of cells to be frozen and what cells are to be used for, but other quantities are OK if necessary.
We also make a test ampoule with most freezedowns. The test contains 1.7 x 10e5 cells/0.17 ml freeze media, or, 1/3 the usual amount of 0.5 ml. They are seeded into 3 35 mm dishes one week after storage of freeze-down in liquid nitrogen to determine the viability and health of the cells in that freezedown.
We feed cells the day before they are to be frozen. If there are multiple dishes-some of which are to be subcultured, we split those one day, and the next day check to see if they are OK (not contaminated). If so, then we go ahead and freeze the remaining dishes.
Follow subculture procedure through #8
9) Bring up (noted) volume of 15 ml or 50 ml test tube to maximal with PBS.
10) Centrifuge the cells in a table top centrifuge at 800-1000 rpm for approximately 5 minutes.
11) While cells are centrifuging, count cells in haemocytometer (10X power). We count 5 squares per chamber x 2 chambers. Record # of cells counted/chamber, and determine total cell count (using the noted volume in the centrifuge tube). Based on what's to be done with cells, calculate volume of freeze medium to be added when cells are resuspended to give 10e6/ml.
12) Carefully aspirate the PBS/STV away from the cell pellet.
13) Bring the cells up to the desired cell density with CPMII. Keep pellet on ice.
14) Label ampoules for freezing (some can be done in advance). Add appropriate amount of cells to each ampoule. Keep ampoules on ice.
15) We use a very low-tech freezing method. It's probably not the best, but it works. Ampoules are wrapped in Kim Wipes and placed in double or triple styrofoam cups lined with crushed Kim Wipes to keep them from freezing too quickly, and then covered with foil.
16) The cup with ampoules is placed in a -80°C freezer for 24 hours.
17) Ampoules are transferred to liquid N2 within one week. (they will start dying after one week at -80°C).
CPMII: stored at -20°C; kept at 4°C after thawing. To make 200 ml: |
||
DMSO |
12 ml |
6% |
FCS |
88 ml |
44% |
DME/F-12 base |
100 ml |
50% |
Mix in T-75 flask. Filter (0.2 µm) for sterility. Do sterility check.
(references: Hammond et al. 1984, Stampfer 1985)
Feeding and Plating Volumes and Densities (for routine culture):
Seeding Cells from a Frozen Ampoule (Non Organoids):
See procedures for cells grown in in serum-containing media. However, MCDB 170 is a clonal medium so it will also support growth of the HMEC seeded at clonal densities. Note: Cells growing serum free are more sensitive to toxins in the environment. Be sure you have not placed in or used for cleaning your incubator any potentially toxic compounds. Washing the incubator with distilled water and then leaving the door open for a day can alleviate some problems.
Cells are frozen in basal MCDB 170 with 10% glycerol plus 15% FCS (no DMSO)(Glycerol I).
Glycerol I: stored at -20°C; kept at 4°C. after thawing. To make 200 ml:
Glycerol |
20 ml |
10% |
FCS |
30 ml |
15% |
MCDB 170 base |
150 ml |
75% |
Mix in T-75 flask. Filter (0.2 µm) for sterility. Do sterility check.
For subculture, see procedures for cells grown in in serum-containing media up to #8. The main difference for cells grown in serum-free media is that the absence of serum in the medium requires the need to pellet the cells to remove all traces of the trypsin.
9) Bring up (noted) volume of 15 ml or 50 ml test tube to maximal with PBS to dilute STV.
10) Centrifuge the cells in a table top centrifuge at 800-1000 rpm for approximately 5 minutes.
11) While cells are centrifuging, count cells in haemocytometer (10X power). We count 5 squares per chamber x 2 chambers. Record # of cells counted/chamber, and determine total cell count (using the noted volume in the centrifuge tube). Based on what's to be done with cells, calculate dilutions to be performed when cells are resuspended. Label dishes/flasks to be seeded, and add necessary amount of medium.
12) Carefully aspirate the PBS/STV away from the cell pellet. Keep pellet on ice.
13) Bring the cells up to the desired cell density with:
For freezing, follow the general instructions for serum-containing media. Cells growing in MCDB 170 are frozen in Glycerol I freeze media. This must be kept on ice during use.
14) Label ampoules for freezing (some can be done in advance). Add appropriate amount of cells to each ampoule. Keep ampoules on ice.
15) We use a very low-tech freezing method. It's probably not the best, but it works. Ampoules are wrapped in Kim Wipes and placed in double or triple styrofoam cups lined with crushed Kim Wipes to keep them from freezing too quickly, and then covered with foil.
16) The cup with ampoules is placed in a -80°C. freezer for 24 hours.
17) Ampoules are transferred to liquid N2 within one week. (they will start dying within one week at -80°C)
(references: Labarge et al. 2013; Stampfer, 1985.)
A. Tissue Processing
1. Obtain human mammary tissue as discard material from surgical procedures, e.g., reduction mammoplasties, mastectomies, biopsies, gynecomastias.
2. Place material in sterile containers containing buffer or media (e.g. 1:1 Ham's F-12 and Dulbecco's Modified Eagle's Medium) supplemented with 10% Fetal Bovine Serum, 4.5g/ml glucose, 100 U/ml penicillin, 100 ug/ml streptomycin, 5 ug/ml Fungizone and 50U/ml polymyxin B and transport to laboratory at 4°C. Reduction mammoplasty tissue can be stored or shipped at 4°C for at least 72 hr without significantly affecting subsequent cell viability. This may not be true of the smaller tissue pieces or tumor tissue.
3. Separate the epithelial areas from the stromal matrix of adipose tissue, connective tissue and blood vessels using a combination of sterile scalpel, forceps and scissors.
a. Nontumor specimens: Transfer cut pieces of tissue into a large sterile dish (e.g.,150 mm). Epithelial areas appear as white strands embedded in the yellower stromal matrix. Gently dissect out these areas, scraping away the grossly fatty material. Lacerate the epithelial tissue using opposing scalpels. Place the epithelial material into a 50 ml test tube. Remove fatty material from the dish for disposal in the autoclave. In heavily fibrous tissues, there will be more solid white, non-epithelial material. We have sometimes encountered difficulty getting much viable cells from such tissues (e.g., subcutaneous mastectomies with severe fibrocystic disease).
b. Tumor Tissue: carefully mince the whole specimen with scalpel and forceps.
4. Place the minced epithelial tissue into a conical centrifuge tube (50 ml or 15 ml) with the tissue comprising no greater than a third of the volume of the tube. Bring the tube up to full volume, leaving only a small air space to allow for mixing during rotation, using a tissue digestion mixture of Ham's F-12 (or equivalent), 10 ug/ml insulin, antibiotics as above, and final concentration of 10% FCS, 200U/ml crude collagenase [Type I, Sigma C3010] and 100U/ml hyaluronidase [Sigma H3506]. Place tubes on a tube rotator and rotate overnight at 37°C.
5. Centrifuge the tubes at 600g for 5 minutes. Discard the supernatant fat and medium. Check for completion of digestion, by diluting a small aliquot of the pellet in medium for microscopic examination. Digestion is complete when microscopic examination shows clumps of cells (organoids) with ductal, alveolar, or ductal-alveolar structures free from attached stroma (see figure P1). Tumor tissue may only show unstructured clumps of epithelial cells. Reduction mammoplasty tissues usually requires additional digestion time. Resuspend the not fully digested pellet in fresh tissue digestion mixture with FCS and enzymes and reincubate with rotation at 37° C for another 4-12 hr or until digestion is completed (occasionally, an additional overnight in fresh digestion mixture may be needed). Use around the same volume of pellet to digestion mixture.
Click here to see Figure P1.
Figure P1: Organoids derived from enzymatic digestion of reduction mammoplasty tissue. The organoid population contains ductal (a), alveolar (b), and ductular-alveolar-like (c) structures freed from the surrounding stroma.
6. When digestion is completed, centrifuge tubes at 600g for 5 min, aspirate the digestion mix (being careful not to disturb pellet), resuspend pellet in medium plus antibiotics at approximately 15 ml/50 ml tube, 5 ml/15 ml tube.
7. Place a sterile 100µ cell strainer (BD Biosciences) on top of a sterile disposable container. Using a Pasteur pipette, place the reconstituted organoid solution on top of the cell strainer, a few ml at a time. Let the medium drain off into the beaker, rewash the organoids with 2-3 ml of medium. Continue the procedure until all the organoids are on the cell strainer. If there are too many organoids and the medium no longer drains away, carefully flip the cell strainer on top of another clean container, wash the organoids down with more medium, then reflip the strainer on the right side on top of the original container and continue the process. Alternatively, or in addition, use a new cell strainer. When completed, flip the cell strainer on top of the collecting container (with organoids facing down) and wash the organoids into the container. This is the 100µ organoids, which contains mostly ductal structures.
8. With a Pasteur pipette, put the filtrate medium from the first container onto a 40µ cell strainer placed on top of another container. Reflip the filter onto another container to wash down the smaller organoids. This is the 40µ organoids that contains mostly alveolar structures. The filtered medium from this second container contains mostly single cells (stromal fibroblasts and some epithelial cells). This is the filtrate. NOTE, the size of the cell strainers used may vary with the material, e.g., for tumor tissues one may start with a 40µ filter. Remember that these methods were developed for normal tissues, and better methods for processing tumor tissues have been developed by others.
9. Transfer the 100µ organoids, 40µ organoids, and filtrate material to 50 ml tubes and centrifuge at 600 g for 5 min.
10. Aspirate the supernatant, reconstitute each tube in CPM I (DME/F-12 or equivalent with 15% FCS and 10% DMSO) using approximately 1 ml of CPM I per 0.1 ml of packed pellet.
11. Seed a test dish for each tube by placing 0.1 ml of resuspended material into 35 mm dish drop by drop to fill in different area of the dish. Let sit for approx. 1 min, then add 1ml of growth medium into dish. Incubate at 37°C and look for attachment the following day. If organoids attached, add another ml of growth medium into dish. Fluid change the dish every other day and watch for growth of organoids as described below.
12. Aliquot the remaining resuspended material into ampoules (1 ml/ampoule). Freeze overnight at -70°C and then transfer promptly to storage in liquid nitrogen. We have not observed any significant loss of viability in our original ampoules stored frozen since the late 1970's.
B. Seeding Frozen Organoids
1. Quickly thaw the frozen ampoule containing the organoids in a 37°C water bath. One ampoule contains approximately 0.1 ml of pelleted organoids in a 1 ml volume of cell preservative medium. Seed the organoids into 2 to 10 (usually around 6) T-25 flasks or 60mm dishes, depending upon visual estimation of the number of organoids in the ampoule.
2. The thawed organoids are carefully placed, drip by drip, onto the surface of the flask or dish with a 1 ml pipette or pasteur pipette for an even distribution of organoids. Avoid scratching the vessel surface (the cells tend not to grow past scratched surfaces). Two ml of growth medium is then added slowly to avoid dislodging the organoids. Incubate at 37°C in humidified CO2 incubator. These are primary culture flasks.
3. After 1 day of attachment, check that the organoids are attached. Add an additional 2ml of medium. Cell outgrowth from the organoids should be visible by 24-48 hours after seeding. (see figure P2A)
4. Feed the cultures regularly 3 times per week. Cells grown in M87A type media grow to near confluence within 5-8 days, while cells grown in MCDB 170 media may take longer (10-14 days).
Differential Trypsinization to remove fibroblasts: Fibroblastic cell growth may be observed in the primary cultures, particularly with tumor specimens or material collected on a filter smaller than 150µm. When the epithelial patches become large, the fibroblasts can be removed by differential trypsinization:
5. Aspirate media. Wash dish/flask with 1-2 ml STV; aspirate. Add 0.5 ml STV (saline, 0.05% trypsin, 0.02% versine) into culture flask at room temperature. Leave the STV on the cells for around 1 minute with continuous microscopic observation to see when the fibroblasts detach but the epithelial cells are still adherent. Knock the dish/flash gently to dislodge the fibroblasts and then quickly aspirate, wash once with PBS, and aspirate again. For cells growing in the serum-free MCDB 170, all traces of the STV must be removed. It is therefore essential that the flask be washed an additional 2 times with PBS before refeeding. M87A grown cells can be refed with medium without additional washes.
C. Subculture of Primary Organoid Cultures
Primary cultures are subcultured when large epithelial patches are present, but before confluence. The density of organoid seeding and attachment will influence the time required. To retain the primary culture and to generate multiple secondary cultures, spaced over time, we perform partial trypsinizations.
1. Aspirate media. Wash with 1-2 ml STV. Add 0.5 ml STV to dish/flask at room temperature. Observe cell detachment under the microscope at room temperature for 1-5 minutes, with gently knocking of the dish to promote cell detachment. The trypsinization should be stopped when around 50% of the cells have detached. Early partial trypsinizations usually have rapid cell detachment. For later partial trypsinizations, cells can be placed at 37°C for faster detachment, but this should be carefully monitored, as all the cells may come off quickly. Trypsinization is stopped as follows:
a) M87A Type Media Cells: Add 2 ml M87A media to the flask, repipette to wash flask and transfer this wash to a 15 ml tube. Repeat with another 2 ml M87A media, again adding the wash to the 15 ml tube. Because the serum in M87A inactivates the trypsin, the primary flask does not have to be washed as is done for the MCDB 170 cells, nor do the cells need to be centrifuged. Refeed the primary flask and return to incubator. Count the cells in the tube with the hemocytometer and seed the dishes directly from the tube to provide second passage cultures. See Procedures "growing and subculturing cells in M87A" for further methods.
b) MCDB 170 Type Media Cells: Add 2 ml PBS to flask; repipette to wash flask and transfer this wash to a 15 ml tube. Repeat with another 2 ml of PBS, again adding this wash to the 15 ml tube. It is essential to wash the primary flask an additional 2 times with PBS to eliminate all traces of STV (discard washes). Refeed the primary flask and reincubate. Note the volume of PBS in the tube, and use a Pasteur pipette to transfer for counting cells in a hemocytometer. Bring the volume of the tube up to 15 ml with PBS and centrifuge at 600 g for 5 minutes. Count the cells in the hemocytometer. Aspirate the supernatant and resuspend the pellet in fresh media. Seeding these cells gives secondary cultures (no longer primaries). See Procedures "growing and subculturing cells in serum-free medium" for further methods. See below for selection of cells in MCDB 170.
2. Partial trypsinization can be repeated around 3 to 12 times per flask, with cell regrowth of the primary flasks and good growth as secondary cultures. MCDB 170 cells need to be subcultured at about 96-hour intervals from first confluence, and M87A cells at about 48-hour intervals. Do not allow the flasks to remain at confluence, or subsequent growth capacity and colony forming ability will be diminished. Removed cells may also be stored frozen instead of seeding. Repeated partial trypsinization of primary cultures in MCDB 170 leads eventually to a selected cell population (see below).
D. Selection of Cells in MCDB 170
1. HMEC from early partial trypsinizations will grow actively, with typical cobblestone morphology, for around 2-3 passages (see figure P2A and P2B). At passage 2-3, most of the cell population gradually changes morphology, becoming larger, flatter, striated, with irregular edges (see figure P2C).
Click here to see Figure P2A, B, and C.
Figure P2: Morphology of normal HMEC grown in MCDB 170. A, primary culture 5 d after seeding. The remnant of the organoid is in the center, with actively dividing cells on the outside of the patch. X38. B, confluent 9th passage cells, with typical epithelial polygonal morphology and continuing mitoses. X152. C, 3rd passage cells. Note the two distinctly different cell morphologies. The smaller cells seem similar to those in (A) and (B). The larger, elongated cells do not maintain miotic activity. X38.
2. When this process of selection first appears, maintain cell cultures with regular feeding, but without subculture. Within a few days to several weeks (usually around 2 weeks) small pockets of actively growing cells with the typical epithelial cobblestone morphology appear (often seemingly "out of the blue", see figure P2C). These cells will push aside the non-dividing large cells and eventually take over the culture, yielded pure post-selection populations.
3. Alternatively, or in addition, my current preferred method of obtaining post-selection cells is through continued partial trypsinization of the primary cultures. After around 5-10 partial trypsinizations, the organoids no longer show widespread vigorous regrowth of actively dividing cobblestone cells. Instead, flat, non-dividing cells predominate. With continued trypsinizations, most organoid areas will eventually give rise to post-selection cells. It may be initially difficult to distinguish between the original good outgrowth cells, and post-selection cells (which are a little more refractile, rounded "perky"-looking). The ultimate test is that non-selected cells will fade in secondary culture, whereas post-selected populations will maintain active growth.
E. NOTES on Propagation of Cells in MM
1. Most reduction mammoplasty derived cultures will show active growth for 3-5 passages in MM. By the 4th or 5th passage, the cell population becomes morphologically heterogeneous, with pockets of actively dividing cells mixed with larger, vacuolated, non-dividing cells (see figure P3A and P3B). In MM type formulations without the conditioned media, we occasionally see some cell growth continuing to higher passage levels.
Click here to see Figure P3A and B.
FigP3: Morphology of normal HMEC grown in MM. A, confluent 2nd passage cells with typical epithelial cobblestone morphology continuing in the confluent populations. (Magnification X 150.) B, confluent 4th passage cells. Note the heterogeneity in cell morphology, with areas of small refractile growing cells, tightly packed areas of small nongrowing cells, and large squamous vacuolated cells. (magnification X 45).
2. Cells grown in MM may produce a lot of lactic acid, which is not as well buffered in media using bicarbonate rather than HEPES buffer bases. We have sometimes needed to remove near-confluent cultures to an incubator with 1-3% CO2 instead of the usual 5%, to avoid levels of acid that are toxic to the cells.
3. Since cells grown in MM have a tendency to multilayer, and cAMP stimulators appear to enhance this property, we usually omit the cholera toxin from the MM formula for growth in primary culture. The cholera toxin is added when cells are placed into secondary cultures.