ES Cell Culture Protocols

Requirements

1. Sacrifice the pregnant mouse by CO₂ asphyxiation.
2. Lie the mouse on its back and swab with 70% ethanol. Using scissors make a cut across the belly and cut away the skin to expose the gut. With sterile forceps and scissors, dissect out the uterus and place it into a petri dish with sterile PBS.
3. Isolate the embryos from the uterus, and release the embryos from the embryonic sacs. Transfer embryos to a second petri dish with sterile PBS.
4. Remove the embryo heads and limbs and scoop out the liver, intestines and heart with a pair of forceps. Transfer the embryo carcass into a sterile 15mL tube with a sufficient volume of Trypsin/EDTA to cover the carcasses. Using scissors finely mince the tissue.
5. Incubate the tissue for 15 minutes at 37°C, then pipette the tissue a few times through a glass Pasteur pipette to dissociate the tissue. Allow the large pieces of cellular debris to settle. Remove the supernatant into a fresh tube and add 50 mL of fibroblast media. Transfer to a 154cm² flask. Plate out about 5 embryos/flask. Incubate at 37°C with 5% CO₂.
6. PMEFs should attach and begin to divide in 1-3 days. After 2 days change the medium which should be very acidic (indicated by the media turning yellow in color).
7. When the flask are confluent, usually in 3-4 days, the cultures are ready for freezing. Freeze cells in 10% DMSO at 2x10⁶ cells/vial (labeled P0).

1. When P4 cells reach confluency they are ready for freezing.

2. For ease of handling process plates in 3-4 batches of about 8 plates at a time.

3. Aspirate media, wash twice with 10mL PBS and trypsinise as usual.

4. Add 2-3mL media to each plate to stop Trypsin.

5. Add ~5mL media to one plate and wash down well to collect cells. Transfer the media to another plate and wash down. Continue transferring and washing down each plate. If the media becomes too thick or frothy, transfer the cells to a 50mL tube and continue with fresh media.

6. Once all the plates are washed and cells harvested transfer the contents to a 50mL tube. Wash down all plates a second time using 5-8mL media to ensure all cells are harvested.

7. Continue for next batch of cells. When all plates are harvested mix the cells well before counting.

8. Count cells by placing 10mL onto a hemocytometer, and counting the number of cells on a 25-grid square. The concentration of cells = number counted x10⁴/mL. Multiply by the volume you have for the total number of cells. Freeze cells at 2x10⁶ per vial.

9. Prior to freezing, g -irradiate cells, 25 GRAY.

10. After irradiation, spin down cells, as usual.

11. Aspirate the media and resuspend cells in the appropriate volume of freezing media to obtain 2x10⁶/mL. Finally, aliquot 1mL of cells into cryovials and freeze immediately at -80°C.

12. Transfer vials to liquid nitrogen storage next day.

ES Cell Culture using ESGRO^®

Two alternative ES media recipes are listed. One method utilizes fetal bovine serum, which may result in rapid growth. However more colonies differentiate after two weeks growth. The serum replacement method (Knockout SR, Gibco BRL, Life Technologies) prevents differentiation very well, but cells grow more slowly. This is especially true if cells have been thawed from serum replacement media.  It is recommended that ES cells be at the lowest possible passage number be used for culturing (between passage number 10-20).

1. Treat 8x10cm tissue culture plates with sterile 0.1% gelatin. Swirl 2-3mLs gelatin to fully cover the plate and let stand for 5-10 minutes. Aspirate the gelatin solution and discard. There is no need to dry the plates following treatment.

2. Thaw a vial containing 1x10⁷ ES cells into 4mL ES media containing ESGRO^® (1000 units/mL) and 4mL FBS. Spin down. Resuspend the cells in 10mL ES media. Plate the ES cells onto the gelatinized plates.

3. Check the cells the next day to determine if fresh media is required (indicated by a change of media color to yellow).

4. Check again the next day, replace with fresh media if required. Once the plate is very crowded and colonies are large (may take 2-3 days but is variable), passage 1 in 2.

5. To passage, have 2x10cm gelatinized plates ready. Remove ES media, wash plates with PBS twice, and add 1.2mL Trypsin. Incubate at 37°C for 2 minutes. Add 10mL ES media and pipette vigorously to break up all the clumps (ES cells tend to stick together which can make counting difficult).

6. Add 5mL to each of two gelantinized plates containing 5mL ES media. If you have 2 plates growing and don’t want to use all the cells, count 10mL on a hemocytometer and freeze the ES cells at between 2x10⁶ and 1x10⁷ per vial. Remember to label the cells with the correct passage number.  ES cells should always be passaged the day before you intend to electroporate.

ES Cell Culture using ESGRO^® and Feeder Cell Layer

1. Treat 8x10cm tissue culture plates with sterile 0.1% gelatin. Swirl 2-3mLs gelatin to fully cover the plate and let stand for 5-10 minutes. Aspirate the gelatin solution and discard. There is no need to dry the plates following treatment.

2. 1 x 10⁶ PMEFs are needed for a feeder layer of this size plate (i.e. half the contents of frozen vial, so 4 vials needed). Thaw PMEFs as usual into 4mL Fibroblast media and 4mL FBS. Spin down the cells and resuspend in 10mL Fibroblast media. Add 5mL of suspension to each of two gelatin coated plates containing 5mL fibroblast media. Incubate the plates overnight to allow the PMEFs to attach and spread.

3. Thaw a vial containing 1x10⁷ ES cells into 4mL ES media containing ESGRO^® (1000 units/mL) and 4mL FBS. Spin down. Resuspend the cells in 10mL ES media. Remove fibroblast media from a feeder plate and plate the ES cells onto the feeder layer. Incubate.

4. Check the cells the next day to determine if fresh media is required (indicated by a change of media color to yellow).

5. Check again the next day, replace with fresh media if required. Once the plate is very crowded and colonies are large (may take 2-3 days but is variable), passage 1 in 2.

6. To passage, have 2x10cm plates of feeder cells ready. Remove ES media, wash plates with PBS twice, and add 1.2mL Trypsin. Incubate at 37°C for 2 minutes. Add 10mL ES media and pipette vigorously to break up all the clumps (ES cells tend to stick together which can make counting difficult).

7. Add 5mL to each of two feeder plates containing 5mL ES media. If you have 2 plates growing and don’t want to use all the cells, count 10mL on a hemocytometer and freeze the ES cells at between 2x10⁶ and 1x10⁷ per vial. Remember to label the cells with the correct passage number.  ES cells should always be passaged the day before you intend to electroporate.

Electroporation

1. On the morning of the electroporation, feed cells with fresh media.

2. Later that afternoon, harvest the ES cells as usual. Count the number of cells in total.

3. 1x10⁷ cells is the minimum number required for electroporation. Calculate the volume of cells needed and reserve. If there is excess, freeze the cells down as previously described.

4. Spin down the volume of cells required for electoporation and remove media.

5. 25-40mg knockout construct DNA (purified) should already be linearized,  ethanol precipitated and dried as a pellet. Dissolve the DNA pellet in 30mL PBS inside the hood. Add to the ES cell pellet (no mixing needed).

6. Add 600mL sterile MT-PBS inside the hood. Transfer to a 0.4cm cuvette and place on ice for 8 minutes.

7. Electroporate at 500mF, 0.24kV (no pulse controller needed). Turn capacitance extender on. A good time constant should be between 8-12ms. Incubate on ice for 10 minutes.

8. Transfer cells to 40mL ES media using a Pasteur pipette. Mix.

9. Transfer 10mL to each of 4 pre-prepared feeder plates (remove the fibroblast media first!).

10. Incubate for approximately 36 hours prior to selection. To select for transformants, add ES media containing between 150-300mg/mL G148 (Geneticin).

11. After 48 hours cell death should be obvious. Feed cells every day if there is a lot of debris and media is yellowing, otherwise every second day is sufficient. If debris is sticking to the living cells, wash the cells gently with PBS before feeding with fresh media. Be careful not to dislodge the feeder layer. Keep growing for around 10 days post electroporation.

Colony Picking

Colonies are generally ready for picking 9-11 days after electroporation, depending upon the cell line and the media used. The best colonies to select are rounded or oval in shape with a phase bright edge, and often a dark necrotic center. Colonies that are differentiated may appear flat, and be surrounded by cobblestone of fibroblast like cells. These colonies are not picked. Medium sized colonies are easiest to pick. Colonies that are considered to be "less than perfect’ are often acceptable, as they tend to bounce back in fresh media after breaking up the colony. Since colonies grown in serum replacement media do not differentiate as readily, they can be grown for longer than 10 days and picked more slowly.

ES cells can be picked onto either gelatinized plates or a feeder cell layer depending upon the ES cell line used. If gelatinized plates are preferred, please disregard the use of feeder cells as described in the procedure below.

1. The day before picking, treat a number of 24 well plates with Gelatin (a few drops). Approximately 5 x 10⁵ PMEFs should be used per plate (i.e. 1 vial can be used on 4x24 well plates). Resuspend PMEFs in 48mL Fibroblast media. Add 0.5mL to each well containing 0.5mL fibroblast media. Incubate as usual.

2. When ready to pick colonies, wear gloves, gown and face mask. Wipe down the microscope, bench, tip boxes and pipette with ethanol. Keep dish closed as much as possible and keep traffic in the room to a minimum.

3. Look at cells at 4x magnification. Any colonies picked should be spaced well enough apart to ensure no contamination from surrounding colonies. When a desired colony is found remove lid, and using a yellow tip and pipette set to 15mL, circle the colony with the tip to loosen the surrounding fibroblast layer (otherwise colony may stick). Scrape the colony with the tip to dislodge, then aspirate in 15mL. The colony can usually be seen inside tip. Transfer to an empty well in a 96 well plate.

4. Keep picking, using and transferring to fresh well using a fresh tip each time, until a suitable number is picked. Clones are often picked in batches of 48 cells to prevent fatigue.

5. Transfer a 96 well plate to the hood. Add a single drop of Trypsin to each well and incubate 37° C, 2 minutes.

6. Replace the media in 24 well plates with 0.5mL ES media containing Geneticin.

7. With a pipette set to 50mL, pipette up and down to break up each colony, careful not to cause too excessive foaming. The better the colony is dispersed the faster it will grow. Transfer to the 24 well plate (transfer any foam also). Use a fresh tip for each well.

8. When all the colonies are transferred, mix each well using a clean blue tip set to 400mL. Incubate as usual.

9. Colonies should start appearing within a few days. If the colonies are few or too close together, disperse them using a blue tip to break up the colonies and spread the cells (Trypsin is not required as colonies break up very easily). Each well should ideally be evenly covered with colonies before harvesting.

10. Continue feeding every second day or whenever the media turns yellow, until a good coverage of colonies in each well is achieved. This usually takes about 7-10 days.

Harvesting and DNA Preparation

1. Remove the media leaving about 500mL in each well.

2. Label an appropriate number of eppendorf tubes to identify each well eg. 1.7A1- electroporation 1, plate 7, well A1.

3. Manually pipette up and down with a blue tip, set to about 400mL to resuspend the cells. Transfer 400mL to each eppendorf tube.

4. When all the wells have been harvested, add 0.5mL fresh media to each well and return to incubator to regrow. Cells should regrow in about 3-5 days. Feed when required.

5. Take the eppendorf tubes to the lab bench to continue. Pellet cells for 30 seconds and remove the media.

6. Add 200mL freshly made lysis buffer. There is no need to resuspend cells. Leave at 37°C in an incubator (not waterbath) overnight.

7. The next day, add 37.5mL 8M Ammonium acetate.

8. Add 250mL Phenol/Chloroform/Isoamyl alcohol in the fume hood. Mix by inversion approximately 5 times. DO NOT VORTEX! Centrifuge 5 minutes.

9. Remove the upper aqueous layer with a pipette tip, leaving any interface behind. If a clumpy clear blob comes up don’t worry. Transfer to 750mL (3 volumes) of 100% Ethanol. Mix well. Often a precipitate is visible immediately.

10. Precipitate at -20°C for 1 hour. Centrifuge 10 minutes, wash with 300mL 70% ethanol and centrifuge again for 5 minutes. Air dry the pellet.

11. Redissolve the pellet in 100mL TE (less if pellet is very small). Allow the pellet to completely dissolve for 2 hours at 65°C then overnight at 4°C.

12. Before digesting, heat to 65°C for 10 minutes. If pipetting is very difficult, pipette the solution straight from 65°C block. Depending upon the size of the original DNA pellet, between 10-30mL of DNA should be digested for Southern analysis.

This method works best with actively growing culture of ES cells (i.e. 1-2 day culture).

Method:

1. The day before, passage a 70% confluent ES cell plate 1:2.
    
2. On the morning of karyotyping, change the medium on the plate at least 3 hours before passage and collection.
    
3. Trypsinize and collect ES cells into a conical tube. Centrifuge as usual and aspirate medium. Avoid allowing the pellet to dry out.
    
4. Gently flick the tube to resuspend the cell pellet, and add 8mL of hypotonic KCl solution to the cells. Continue to gently flick the tube during the addition of KCl to avoid clumping.
    
5. Incubate the tube at 37°C for 10 minutes (this may vary for each type of cell line used).
    
6. Add 2mL of freshly made fixative and mix by gentle inversion.  {Fixative MeOH:Glacial Acetic acid 3:1 made fresh and stored at 4°C}.
    
7. Centrifuge cells at 1000 rpm for 5 minutes and aspirate supernatant.
    
8. Using a pasteur pipette, carefully add 2mL of fixative solution dropwise, with gentle mixing to avoid clumping. Add an additional 6mL of fixative and mix by gentle inversion of the tube.
    
9. Centrifuge cells at 1000 rpm for 5 minutes and aspirate supernatant.
    
10. Repeat steps 8 & 9 three times.
     
11. Resuspend the pellet in 1mL of fixative (less or more according to pellet size).
     

To make cell spreads, firstly humidify the surface of a dried cold slide by application of warm breath, whilst holding the slide at a 45° angle. Using a pasteur pipette, carefully drop (from a height of approx 0.5 metres) one drop of the suspended cells onto the top surface of the slide and allow to air dry.

Staining:

1. Stain slides with freshly made Leishmann’s stain for 8 minutes.
    
2. Rinse in running water for 1 minute and air dry.
    
3. Clear cells in 2x changes of xylene and mount coverslip using Depex.

Notes:

Leishmann’s Stain (BDH Product #35022)
Make to 0.2% w/v solution in methanol.

ES Media (with Fetal Bovine Serum):
200mL fibroblast media
2mL non-essential amino acids
ESGRO^® (1000 units/mL is recommended).
For selection media add:
150-300mg/mL G418 (Geneticin)

ES Media with Serum Replacement:
500mL knockout DMEM (GibcoBRL, Life Technologies)
90mL knockout serum replacement (15%) (Knockout SR, GibcoBRL, Life Technologies)
6mL 100x non-essential amino acids (Trace Biosciences Pty. Ltd)
6mL 100x pen/strep
6mL 100x glutamine
4.6mL mercaptoethanol
ESGRO^® (1000 units/mL is recommended).

MT-PBS:
0.227g Na₂HPO₄
0.0625g NaH₂PO₄.2H₂0
0.87g NaCl

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