Procedures for Culture of Human Mammary Epithelial Cells


Table of Contents
  • I. Cells Grown In a Serum Free Medium


  • I. Cells grown in serum free medium.

    (references: Hammond et al. 1984, Stampfer 1985)

    A. Making media and stocks.

    We obtain our media, originally designated MCDB 170, from CLONETICS (1-800-85CLONE;); other sources of MCDB 170 are available.

    The media from Clonetics comes in various forms with different names. Their version of complete MCDB 170 with most of the serum-free supplements is called MEGM, Mammary Epithelial Growth Media. NOTE: MEGM does not contain all the serum-free supplements that we use, i.e., it does not contain transferrin and isoproterenol. These need to be added. Also, the BPE is supplied separately. Clonetics also provides a medium lacking most of the serum-free supplements, which is called MEBM, Mammary Epithelial Basal Medium. NOTE: this medium does contain the ethanolamine and phosphoethanolamine - do not add these. Additionally, the media can use either a sodium-bicarbonate buffer base, or a HEPES buffer base, called MEBM-SBF. We highly recommend you use the HEPES based (sodium-bicarbonate free) media if at all feasible. Clonetics also provides a phenol-red free medium, called MEBM-PRF. The serum-free supplements that need to be added to each 500 ml bottle (if not already present) are as follows:

    Supplement Amount/Bottle Final Concentration
    EGF (20,000X stock, 100ug/ml) 25 ul 5 ng/ml
    Hydrocortisone (2000X stock, 1 mg/ml) 0.25 ml 0.5 ug/ml
    Insulin (200X stock, 1 mg/ml) 2.5 ml 5 ug/ml
    BPE (200X stock, 14 mg/ml) 2.5 ml 70 ug/ml
    Transferrin (2000X stock, 10 mg/ml) 0.25 ml 5 ug/ml
    Isoproterenol (500X stock, 5x10^-3 M ) 1.0 ml 10^-5 M
    Ethanolamine (1000X stock, 10^-1 M) 0.5 ml 10^-4
    O-Phosphothanolamine (1000X stock, 10^-1M) 0.5 ml 10^-4M

    Maintenance of the appropriate pH is critically important. We have kept cells grown in the original formulation of MCDB 170 (HEPES based) in incubators with low (0.1-2%) CO2 settings. At these low settings there may be considerable variation among incubators. Ultimately, proper pH is best determined by color indicator: media should be salmon orange. Yellow is too acid, red is too basic. Clonetics media with bicarbonate is designed to make it easier for people to use the cells at the more typical setting of 5% CO2. However, the cells do not grow as well with the bicarbonate base, do not maintain their pH levels as well inside or outside the incubator, and can be less efficient for certain procedures (e.g., calcium phosphate mediated transfection). I strongly recommend that you obtain the HEPES buffered media if you can designate an incubator to be used at the low CO2 setting. 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.

    MCDB 170 is also available from the UCSF Cell Culture facility (415 476-1450). While this media is cheaper, it requires more effort to put together (check with me for instructions) and there is significantly less quality control (e.g., the media is not tested on the mammary cells). Other commercial sources provide powdered MCDB 170.

    Serum-free Supplement Stocks for MCDB 170

    BPE (Bovine Pituitary Extract) 200x stock, 14 mg/ml

    We obtain our BPE from Susan Hammond, 510 522-3625, P.O.Box 147, Alameda, CA 94501. This is the cheapest source we know but requires some processing. BPE is also available ready to use from Clonetics and other commercial sources.

    1. Take bottles (100 ml) "raw" BPE out of freezer and put in 37 C water bath to warm up.
    2. Transfer BPE to several Oak Ridge tubes and balance pairs.
    3. Centrifuge using adaptors in JA-20 rotor at 15K rpm for 30 min., 4 C . Make sure that the "O" rings on the rotor are in place.
    4. Pour the supernatant into clean Oak Ridge tubes. Rebalance. Centrifuge again at 15K for 10 min. more. If supernatant doesn't appear to be clear, centrifuge 10 min. more, same conditions.
    5. Transfer supernatant from all tubes into flask. Discard the pellet at the bottom of the tubes. Filter the supernatant through 0.8 µm, 250 ml Nalgene filters, about 30 ml at a time; then filter through 0.2 µm, 115 ml Nalgene filters. Several will be needed because they get clogged.
    6. Label sterile 50-ml and snap-top polypropylene tubes with "BPE, 200X, month/year, vial letter, i.e. A, B, C, etc." and include an expiration date.
    7. Aliquot 26 ml and 2.8 ml portions into sterile polypropylene tubes.
    8. Do sterility check on each vial, using 10 µl from each aliquot into 1.5 ml sterile media, check test every day for 4 days.
    9. Store in labeled rack in -20 °C

    EGF (Epidermal Growth Factor) 20,000x stock, 100 µg/ml

    We obtain Human EGF from Upstate Biotechnology. Many other commercial sources are available

    1. Retrieve 100 µg vial of hEGF from refrigerator. Make sure that the vial is unopened and actually says 100 µg.
    2. Make up to 0.1 mg/ml by adding 1.0 ml sterile distilled water to vial. Mix gently, but well. If necessary, vary the concentration according to the weight in the vial.
    3. Aliquot 0.26 ml portions into sterile ampoules labeled "hEGF, 20,000X, Month/Year, ampoule letter, i.e. A,B,C etc."
    4. Check sterility of each ampoule by adding 3 µl from each ampoule to 1.5 ml media in a 35 mm dish and incubate 3 or 4 days. Check every day for contamination.
    5. Store in -20 °C freezer for up to 3 months

    Transferrin Human 2000x stock, 10 mg/ml We obtain from Sigma CAT# T-2252 Siderophilin

    1. Dissolve 1000 mg of transferrin into 100 ml distilled water. This gives a stock concentration of 10 mg/ml.
    2. Filter for sterility through 0.2 µm filter.
    3. Aliquot 2.6 ml and 0.30 ml portions into sterile polypropylene tubes or snap-top tubes labelled "Transferrin, 2000x, month/year, vial letter, i.e., A,B,C, etc."
    4. Do sterility check by placing 10 µl from each vial into corresponding labeled 35 mm dish with 1.5 ml media, check every day for 4 days.
    5. Store at -20 °C freezer.

    Isoproterenol 500x stock, 5x10-3M We obtain powder from Sigma CAT #: I-5627 (+ Isoproternol: hydrochloride crystalline) which is stored at room temperature.

    1. Make up in hood. Do not inhale dust. Measure in hood or in covered balance. To make up 40 ml, use 50 mg IP in 40 ml 95% EtOH.
    2. Store stock in freezer: Make up fresh monthly.

    Insulin 200x stock, 1 mg/ml We obtain from Sigma CAT # I-5500

    1. Dissolve 1g of powder in 200 ml of 0.005 N HCl (1 ml 1 N HCl with 199 ml of distilled water) by stirring on a magnetic stirrer.
    2. When the solution is clear*, add 800 ml of distilled water, to make the final concentration of insulin 1 mg/ ml.
    3. Sterilize by filtering through a 0.2 µm filter.
    4. Label approximately 30 sterile snap-top tubes and enough 50-ml sterile polypropylene tubes with "Insulin, 200X, month/year". Aliquot 2.8 ml and 26 ml portions into sterile polypropylene tubes.
    5. Store at -20 °C.

    * If the solution is not clear after a reasonable amount of stirring, add a few more drops of 1 N HCl. (The total [HCl] should not exceed 0.005 N HCl/ liter of solution.) When the solution clears, then bring up to 1 liter with distilled water.

    Hydrocortisone 2000x stock, 1 mg/ml We obtain from Sigma CAT #H-4001

    1. Add 50 mg Hydrocortisone to 50 ml 95% ethanol, mix well.
    2. Store at -20 °C in a sterile 100 ml glass bottle, no sterility test necessary.

    Ethanolamine (2-Aminoethanol) 1000x stock, 10- 1M We obtain from Sigma: CAT# A-5629

    1 Dissolve 341.6 mg ethanolamine (approx 0.15 ml) in 56 ml MCDB 170 base to give a final stock concentration of 6.1mg/ml or 0.1M.
    2. Filter through 0.2 µm filters for sterility.
    3. Label sterile polypropylene tubes "Ethanoloamine, 1000X, mo./yr, vial letter" and aliquot 5 ml and 0.6 ml portions.
    4. Do sterility test by aliquoting 5µl from each tube into corresponding labelled 35mm dish with 1.5 ml media. Check every day for four days.
    5. Store at -20 °C.

    O-Phosphoethanolamine 1000x stock, 10- 1M We obtain from Sigma CAT# P-0503 .

    1. Dissolve 789.5 mg in 56 ml of MCDB 170 base to give a stock concentration of 14.1 mg/ml or 0.1M.
    2. Filter through 0.2µ filter for sterility.
    3. Label sterile polypropylene tubes "phospoethanoloamine, 1000X, mo./yr, vial letter" and aliquot 5 ml and 0.6 ml portions.
    4. Do sterility test by aliquoting 10µl from each tube into corresponding labelled 35mm dish with 1.5 ml media. Check every day for four days, repeat if necessary.
    5. Store at -20 °C.

    NOTE: Also spelled 'O-Phosphoroethanolamine' and 'O-Phosphorylethanolamine

    B. Growing and Subculturing Cells in Serum-free Medium

    1. Feeding and Plating Volumes and Densities (for routine culture):

    Dish/Flask Amount Media (ml) Number of Cells for Seeding
    T-75 12-15 3-5 X 10^-5
    100mm 10-12 3-4 X 10^-5
    60mm; T-25 3.5-5 1.0-1.5 X10^-5
    35mm 1.5 0.5 X 10^-5
    24 Well Plate 1.0/well 2 X 10^-4/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.

    2. Seeding Cells from a Frozen Ampoule (Non Organoids):

    1) Label and add medium to all dishes
    2) Put 2 scoops of liquid nitrogen 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, wipe down with 70% ethanol in the hood.
    5) Open amp and mix well with added media for a good distribution into the dishes, i.e., if seeding 5x10^-5 cells into 3-60's, assume that there is 0.5 ml in the amp already and add 1.3 ml media, mix and distribute 0.6 ml to each 60 mm dish. It's most important to mix cells well and distribute cells evenly.
    6) Distribute cells evenly by gently pushing dish back and forth 8-10 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 10% glycerol plus 15% FCS (no DMSO).

    3. Subculture of Cells

    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 subconfluent and still actively proliferating. When the cells are growing as they should, they are subcultured around once a week 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. Stor e 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 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) Take cells out of incubator and check under microscope to see if they have all "rounded up". Don't leave the STV on longer than necessary to remove most of the cells - trypsin chews up cells. Tap dish lightly against hand or desk if necessary to kn ock cells loose. Continue incubation at 37 °C. if cells are still attached.
    6) When most of the cells have loosened add PBS 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, or in the case of 184B5, which tends to hold on tightly. Repipette the PBS 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 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) Bring up 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.
    13) Bring the cells up to the desired cell density with: 4. Freezing Cells for Liquid Nitrogen Storage

    We always freeze our cells at the density of 10^-6 cells/ml freeze medium. Cells are generally frozen in 0.5 or 1.0 ml amounts (5 X 10^5 or 10^6 cells), depending on number of cells to be frozen and what cells are to be used for, but other quantities are O K if necessary.

    We also make a test ampoules with every freezedown. The test contains 1.7 X 10^5 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 remai ning dishes.

    1) Follow subculture procedure to # 12
    2) Cells growing in MCDB 170 are frozen in Glycerol I freeze media. This must be kept on ice during use.

    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.

    3) Add the appropriate amount of Gly I to cell pellet to have 10^-6/cells/ml. Keep on ice.
    4) Label ampoules for freezing. Add appropriate amount of cells to each ampoule. Keep ampoules on ice.
    5) 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 a styrofoam cup lined with crushed Kim Wipes to keep them from freezing too quickly, and then covered with foil.
    6) The cup with ampoules is placed in a -70 °C. freezer for 24 hours.
    7) Ampoules are transferred to liquid N2 within one week.

    II. Cells grown in serum-containing MM medium.

    (references: Stampfer et al., 1980; Stampfer, 1982; Stampfer 1985)

    A. Making media and stocks

    The composition of MM Medium as originally formulated is as follows:

    Component Amount/500 ml Final Concentration
    1:1 mixture of DME/F-12 300 ml
    Hs767Bl and/or fHs74Int (**) 100 20%
    Hs578Bst (**) 50 10%
    FCS 2.5 ml 0.5%
    Insulin (100x stock; 1 mg/ml) 5 ml 10 µg/ml
    EGF (20,000x stock; 10 µg/ml) 25 µl 5 ng/ml
    Hydrocortisone (10,000x stock; 1 mg/ml) 50 µl 0.1 µg/ml
    Cholera toxin (10,000x stock; 10 µg/ml) 50 µl 1 ng/ml
    T3;triiodothyronine (20,000 stock; 2x10-4M) 25 µl 10^8 M
    E2;-estradiol (20,000x stock;2x10-5 M ) 25 µl 10^9 M
    Pen-Strep (100x stock)(optional) 5 ml
    ** Conditioned Media: (Cells grown in 1:1 DME/F-12 with 5% FCS + 5 µg/ml insulin)

    The above represents MM medium as originally formulated. We have used many variations on this (unfortunately not all given clearly distinct names). Nowadays, we often omit the T3 and E2 from the formulation. We do not have good experimentation to indicate the effect of these components.

    MM without any conditioned media (CM) is referred to as MM4. The cells will proliferate without the CM as long as the cholera toxin is present. The CM can effect both the long-term growth of the cells and their state of differentiation. Nowadays we often use only Hs767Bl CM (we have frozen stocks of these cells). Early experiments suggested that the CM from Hs578Bst served mainly to help cell attachment.

    The DME/F-12 base is buffered with sodium bicarbonate, generally requiring around 5% CO2 settings for the incubator. However, we have found that actively growing cultures, particularly as they approach confluence, tend to overwhelm the acidic buffering capacity. This can be seen by the phenol red indicator - culture medium becomes yellow. Therefore, we often kept a separate incubator set at lower CO2 concentration (around 3%) to which we removed cells as they became more confluent. Frequent refeeding could also alleviate the acidity. The cells will lose viability if left acid for too long.

    Stocks for MM

    (not already listed for MCDB 170)

    Cholera Toxin 10,000x stock (10 µg/ml) We obtain from Sigma CAT# C3012

    1.Add 0.5 ml sterile distilled water to 0.5 mg of cholera toxin in bottle to yield 1 mg/ml.
    2. Dilute this 1:100 (0.1 ml/10 ml) in sterile distilled water to yield 10 µg/ml. Leave in concentrated form because it loses activity less quickly.
    3. Aliquot into 1 ml ampoules.
    4. Store in refrigerator for up to 3 months; do not freeze.

    T3; 3,3', 5-tri-iodo-L-thyronine 20,000x stock (2x^10-4M) We obtain from Sigma CAT# T-2877.

    1. Dissolve 1.302 mg in 10 ml distilled water. Add 1 drop weak NaOH (0.1 M to 0.5 M). Filter (0.2 µm) sterilize. Refrigerate. Make up fresh monthly.

    E2; beta-estradiol 20,000x stock (2x10^5 M) We obtain from Sigma CAT# E-8875

    1. Dissolve 5.44 mg in 10ml absolute EtOH to make 2 x 10-3M. (MW = 272.4).
    2. Dilute 100 µl of this into 10 ml for a 2x 10-5 M stock. Store in freezer at -20 °C.

    B. Growing and Subculturing Cells in MM

    1. Feeding and Plating Volumes and Densities:

    Feeding and plating densities are similar to cells in MCDB 170 with one major exception. MM does not support clonal growth of HMEC. Seeding at densities lower than those indicated will lead to reduced cell viability.

    2. Seeding Cells from a Frozen Ampoule

    The procedure is the same as for cells grown in MCDB 170 except that the cells grown in MM have been frozen in 1:1 DME/F-12 with 15% FCS and 10% DMSO.

    3. Subculture of Cells

    See general notes for cells grown in MCDB 170. The main difference for cells grown in MM is that the presence of serum in the medium eliminates the need to pellet the cells to remove all traces of the trypsin.

    1) Look at cells under microscope. Make sure they are not contaminated and look appropriate.
    2) Aspirate media from parent dish. Wash once with STV (not something else):

    3) Add enough STV to barely cover the cells, in general:
    4) Incubate cells in 37C. 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) Take cells out of incubator and check under microscope to see if they have all "rounded up". Don't leave the STV on longer than necessary to remove most of the cells - trypsin chews up cells. Tap dish lightly against hand or desk if necessary to kn ock cells loose. Continue incubation at 37C. if cells are still attached.
    6) When most of the cells have loosened add MM to culture vessel, preferably using a plugged Pasteur pipette with a hand pipetter, or otherwise a regular pipette. Do not wait for all the cells to come off, particularly if only small patches remain. Repipette the MM in the flask/dish to break up clumps, and then transfer this to 15 or 50 ml tubes. Wash the flask/dish with about the same amount of MM one or two times and transfer to tube. The final volume in the tube to optimize cell counts 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) 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. Label dishes/flasks to be seeded, and add necessary amount of medium.
    9) Seed dishes and gently swirl medium to insure even distribution of cells in the culture vessel. Place in incubator.

    4. Freezing Cells for Liquid Nitrogen Storage

    See notes for cells grown in MCDB 170. Follow the same procedures (including the subculture procedure for cells grown in MCDB 170). The only difference is that we have used a different cell preservative medium (CPMI) for cells grown in MM. Additionally, it is not necessary to use just PBS to stop the trypsinization. MM, or buffer/medium containing FCS can be used.

    CPMI : stored at -20oC; kept at 4oC after thawing. To make 200 ml:
    DMSO 20 ml 10%
    FCS 30 ml 15%
    DME/F-12 base 150 ml 75%
    Mix in T-75 flask.


    II. Tissue Processing Protocol


    (references: Stampfer, J. Tiss. Cult. Meth. 9: 107-115, 1985.
    or Taylor-Papadimitriou, J., and Stampfer, M.R., Culture of Human Mammary Epithelial Cells. In: Cell & Tissue Culture: Laboratory Procedures, (Griffiths, J.B., Doyle, A., Newell, D.G., eds.), Wiley-Liss, pp107-133, 1992)

    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 pieces of 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-alveolarlike (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 150µ filter holder (filter cloth secured between two metal plate holders) on top of a sterile disposable container. Wet one side of the filter cloth with about 1 ml of medium, and turn the filter over to the other side. Using a Pasteur pipette, place the reconstituted organoid solution on top of the wetted filter cloth, a few ml at a time. Let the medium drain off into the beaker, rewash the organoids with 2-3 ml of medium (by placing medium directly on top of the filter cloth). Continue the procedure until all the organoids are on the filter cloth. If there are too many organoids on the cloth and the medium no longer drains away, carefully flip the filter cloth on top of another clean container, wash the organoids down with more medium, then reflip the filter cloth on the right side on top of the original container and continue the process. Alternatively, or in addition, use a new filter. When completed, flip the filter cloth on top of the collecting container (with organoids facing down) and wash the organoids into the container. This is the 150µ organoids which contains mostly ductal structures.

    8. With a Pasteur pipette, put the filtrate medium from the first container onto a 51µ c filter cloth (treated previously as the 150µ c filter) placed on top of another container. Reflip the filter onto another container to wash down the smaller organoids. This is the 51µ organoids which 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 filter clothes used may vary with the material, e.g., for tumor tissues one may start with a 95µ or 51µ filter. Remember that these methods were developed for normal tissues, and better methods for processing tumor tissues are being developed by others (e.g., see Investigator list for Steve Ethier, Shanaz Dairkee, Vimla Band).

    9. Transfer the 150µ c organoids, 51µ c 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 nunc type 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 disloging 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 outgroth 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 MM 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. MM 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)MCDB 170 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.

    b)MM Type Media Cells: Add 2 ml MM media to the flask, repipette to wash flask and transfer this wash to a 15 ml tube. Repeat with another 2 ml MM media, again adding the wash to the 15 ml tube. Because the serum in MM 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 MM" for further methods.

    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 MM 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.

    Email:mrstampfer@lbl.gov Index Back