Media and Solutions Required for Routine ES Cell Culture
Media used to prepare 100 mL
Routine Culturing of ES Cells
- Dulbecco’s Modification of Eagles Medium (DMEM)
- 1X without L-glutamine with 4.5 g/L glucose. Store at 4 °C
- Cytosystems 500 mL Cat. No. 11.016.0500V
- Nonessential amino acids 100X
- Cytosystems 100 mL Cat No. 21-145-0100V. Store at 4 °C
- Penicillin/streptomycin (5000 1U/mL, 5000 µg/mL)
- Cytosystems 100 mL Cat No. 21-140-0100V. Store at –20 °C
- Fetal calf serum (FCS)
needs to be tested for the ability to support growth of ES Cells. Serum
should be stored frozen and heat-inactivated before use by heating at 56
30 mins. It may be stored at 4 °C following inactivation. The heat
inactivation removes complement activity which, along with natural
antibodies, may be toxic for ES cells.
- 100X nucleoside stock
- To prepare 100 mL (100X)
to 100 mL Travenol water and dissolve by warming to 37°C.
aliquot while warm. Store at 4°C for months. The nucleosides
come out of solution. Warm to 37°C before use to resolubilize. Solution
to be stable at 4°C.
- Other solutions required.
- 1X Dulbecco’s phosphate-buffered salt solution (PBS) without calcium and magnesium.
- Trypsin 2.5% Cytosystems 100 mL 21-159-0100V
- Trypsin/EDTA (1:250) Cytosystems (0.05% Trypsin) 100 mL 21-160-0100V
- Solutions to be made up
- PBS/EGTA (0.5 mM)
mL of 0.05M stock EGTA (Sigma E4378) to 100 mL of PBS, to
produce a final concentration of 0.5 mM.
- Stock EGTA in H2O 0.05M add concentrated NaOH to dissolve
- Trypsin 0.25%.
- Add 1.6 mL 2.5% Trypsin to 20 mL Cytosystems Trypsin/EDTA 0.05%.
- 0.1% gelatin in PBS
- Add 0.1 gm Gelatin Sigma G-1890 and autoclave to 100 mL PBS
- 0.1M 2-mercaptoethanol (§ME) sigma M-6350.
- Add 0.1 mL §ME (14.4M) to 14.3 mL PBS and filter through a 0.2 µM
- ACRODISC. Store at –20 °C for up to 1 month.
- LIF (if no feeders are used)
- LIF ESGRO AMRAD (murine Lif in PBS/BSA solution) 1 mL ampoule
- 107 U/mL. Dilute 1/100 in DMEM 10% FCS, aliquot into 10 × 10 mL
tubes, store at –20 °C until use.
- 107 U/mL. Dilute 1/100 = 105 U/mL. (100X conc)
- Dilute 1/100 for use = 1000 U/mL.
are normally passaged every 2-3 days; this is important to avoid
Signs of differentiation are:
Cells are passaged as follows:
- Colonies are surrounded by flattened, differentiated cells.
- Large colonies with necrotic centers, these appear as cells with defined
- Colonies appear as individual cells rather than as a syncial mass.
are more “rounded” than “flat”, they also have a clearly defined
boundary. Worse than this, they have formed free-floating embryoid
Isolation of Primary Mouse Embryo Fibroblasts
- All reagents are warmed to 37 °C.
Medium: PBS, PBS/EGTA, TRYPSIN/EDTA
- Remove medium.
- Wash with PBS (5 mL/25 cm2 flask, 10 mL/80 cm2 flask), aspirate.
- Wash with PBS/EGTA, aspirate.
- Place flask on 37 °C warming tray for approx. 1 min or until individual
cells can be seen in colonies.
- Add 0.5 mL (25 cm2), 1 mL (80 cm2)
TRYPSIN/EDTA, and rock flask
backwards and forwards until colonies float off; this should take ~1
- Using a 1-mL pipette, pipette up and down, (avoid making bubbles as
kill the cells), for approx. 1 min. Check that all colonies have been
dispersed and that a single-cell suspension has been achieved. Don’t
in TRYPSIN/EDTA for longer than 3 mins, as it is quite toxic.
- Neutralize trypsin by adding an equal volume of medium, mix by gentle
- Aspirate media from feeder flask, as this is different media, to ES cell
media. Seed feeder* flasks with an aliquot of cell suspension. A 1:10
split is appropriate for a well-growing culture with medium–large
colonies that are not touching each other but are reasonably close
*Feeder flasks contain Mitomycin C-treated (see protocol) Primary Mouse
Embryo Fibroblasts (PMEFs) at a concentration of 0.3 × 106/25 cm2 flask,
1 × 106/80 cm2 flask, or Mitomycin C-treated STO cells at a concentration
of 1.25 × 106/25 cm2 flask× 106/80 cm2 flask. STO cells are smaller
cells are to be grown in the presence of LIF only, i.e., no feeder
flasks or plates must be treated with 0.1% gelatin in PBS at 37 °C for
1–2 hrs. This is removed before the medium is added..
- You will
need a 13.5-day pregnant mouse (we use MTK NEO inbred white
- 2 sets of sterile instruments, one containing a pair of curved forceps and
a pair of iris scissors
containing 2 pairs of curved forceps, 1 pair of iris scissors, and a #3
- Phosphate buffer saline (PBS)
- Sterile medium-size petri dishes (tissue culture standard)
- 18-gauge needle
- Luer lock syringe (about 6 cc should suffice)
- #11 size flat-edged scaple blade
- Dulbecco’s Modification of Eagles Medium (with 10% fetal calf serum, 1%
penicillin/streptomycin, 1% L-glutamine, 0.2% 0.1 m BME)
- Large flasks (tissue-cultured standard, about 154 cm2 area)
- Class II Laminar Flow hood.
starting, pour out 2X petri dishes of PBS in the hood. The pregnant
is killed by cervical dislocation. (This is not done in the hood but on
benchcote). Lay mouse out on its back and swab belly with 70% ethanol.
pair of scissors (not sterile), nip a small cut across the belly.
skin above and below the nip with your finger, tear the skin apart and
over the head and hind legs to expose the viscera of the gut. This
cleaner than cutting through the fur and enables you to reach the uterus
of touching the fur (cutting through dry fur creates a bacterial
sterile forceps and iris scissors, dissect out the uterus, taking care
touch the fur or the benchcote with the uterus or instruments. Place the
into a petri dish of sterile PBS and swirl around to remove blood.
Transfer the uterus to a second petri dish of sterile PBS and move the
the second set of sterile instruments and a fresh sterile petri dish,
the embryos. Be sure to remove the placement and embryonic sacs. Using
scalpel handle with the #11 blade on it, cut off the embryo’s head and
liver with a pair of forceps. The head and forelimb should be cut off
the head and liver and leave the bodies in fresh PBS in a fresh
dish. Take a 6-cc luer lock syringe with an 18-gauge needle attached and
the plunger. Keep the plunger sterile. Drop the embryo bodies inside the
and add 3 mL trypsin/EDTA. Put plunger back in syringe and squirt
contents of syringe into a large tissue culture flask. Place the flask
tray (37 °C) for 2–3 minutes. Then place it back in the hood and add
DMEM. When adding the DMEM, try to wash any tissue off the walls
tissue. Transfer flask to an incubator at 37 °C with 5% CO2
. Do not put
less than 7 embryos in one large flask.
Loosen the lid of the flask in incubator to allow gas exchange
in the medium. This is the primary isolation or passage one.
attach and begin to divide in 1–3 days. During this time, do not
disturb, so as
to allow PMEFs to settle and attach. After 2 days, change the
medium. It will
be very acidic. After 3–4 days, the culture will need splitting.
and gently wash the monolayer with 2X 10 mL PBS. Add 2 mL
trypsin EDTA and split 1:4. After a further 2–4 days, the culture will
be ready for freezing.
number obtained from each flask will be between 5–10 × 106 cells. Freeze
in 10% DMSO at 3X 106/ampule.
recovering the cells from LN2, put all the cells into a medium flask.
confluent these are split into 1 medium and 1 large flask (1:3). The
can be treated with mitomycin C and the medium flask split again. Do not
Media for Embryo Culture and Manipulation
Medium (Protocol obtained from Karen Austen-Reed from SS Tan Laboratory,
oocyte maturation and routine culture of mouse embryos, M16 culture
is used. This medium is unable to maintain its own pH, and must
therefore by used in conjuction with an incubator buffered with 5% CO2
maintains the required pH level of the medium.
- Penicillin/streptomycin—use at 1/100 (Tissue culture Pen/Strep)
- Phenol red 0.005
- NaHCO3 25.0 84.02 1.051
- C Na pyruvate 0.33 110.0 0.018
- D CaCl2-2H2O 1.71 147.2 0.126
- BSA (bovine serum albumin) 4 mg/mL
DMEM with HEPES
- Weigh out all of stock “A” (except Pen/Strep and Na lactate) into a
measuring cylinder and make up to 90 mL with “Travenol” water. Add
Pen/Strep to cylinder and then Na lactate (Note: the Na lactate is quite
viscous—by heating it up to ~37 °C prior to use, it can be more easily
accurately pipetted). Make up to 100 mL with “Travenol” water and pour
- Weigh out stock “B” (Phenol red, NaHCO3)
components into a measuring
cylinder. Make up to 100 mL with “Travenol” and pour into a 500-mL
bottle. Weigh stock “C” (Na Pyruvate) into measuring cylinders, make up
100 mL with “Travenol” water and pour it into a 500-mL bottle. Weigh
“D” (CaCl2-2H2O) into a measuring cylinder, make
up to 100 mL
“Travenol” water, and pour it into a 500-mL bottle. Add about 100
“Travenol” water to bring the total volume to 500 mL. Be sure that
components of each stock have dissolved properly before adding them
the 500-mL bottle. Also be sure to add stocks in their alphabetical
avoid precipitation of some ingredients.
- Make 50-mL aliquots of this solution and freeze them. Use one 50-mL
aliquot at a time by aliquoting it into tubes of 9 mL each, then
use. The osmolarity should be 288-292 m osmol.
Before use, lightly gas with CO2,
then add 1 mL of FCS (fetal calf serum)
9-mL aliquot of M16. Then sprinkle 36 mg of BSA (4 mg/mL M16) on
and allow to dissolve—do not shake up or stir. Then Filter-sterilize
mix using a 12-mL syringe with an acrodisc.
Collection of Morulae and Earlier
- This medium is used for manipulations that are performed on the mouse
embryos while out of the incubator. Since there is not 5% CO2 present to
maintain the pH, this medium contains HEPES to keep the pH
- Ingredient % of final volume
- 1 X DMEM with 20 mM HEPES buffer 80.8%
- Penicillin/streptomycin (5000 i.u/mL/5000 µg/mL) 1%
- L-glutamine (200 mM, 100X) 1%
- Nonessential amino acid (100X) 1%
- Nucleosides (100X) 1%
- β-mercaptoethanol (0.1M) 0.2%
- Fetal calf serum 15%
recipe is made up fresh each week (the same stocks of ingredients
be used). An alternative to DMEM with HEPES is M2, the recipe for
can be found in Bridgette Hogan’s Book, “Manipulating the Mouse
- Before you start, have ready the following:
- Sterilize 2 pairs of curved forceps, 1 pair of iris scissors, and 2 pairs
of fine watchmaker forceps.
6-mL syringe filled with DMEM with HEPES, with an 18-gauge
needle attached. To the needle attach a 20-cm length of clear vinyl
into the end of this tubing insert a sharp flusher.
2.5 days pc (post-coitus), the morulae are present in the oviducts. For
reason, it is only necessary to remove the oviducts from the mouse.
the 2.5-days pregnant mouse and lay it on its back on benchcloth or
absorbent paper. Swab its belly with ethanol (70%) and nick the skin
pair of scissors. Pull the skin back over the head and toward the tail.
pull back the body wall to expose the contents of the abdomen. Push
the guts to expose the reproductive tract. Using a pair of curved
forceps, grasp the uterus just below the oviduct and carefully separate
oviduct from the ovary using the iris scissors. Cut through the uterus
the uterotubal junction and place the oviduct in a petri dish.
the dish to the stage of a stereo dissecting microscope. Using the
watchmaker forceps, manipulate the oviduct so that you have the end of
oviduct (close as possible to the uterotubal junction) between the
insert the flusher so that it points away from the uterotubal junction.
careful not to pierce through both sides of the oviduct tube. Keeping
flusher inserted into the oviduct, pick up the syringe in the other
give a couple of short, sharp squeezes on the plunger. This should
all morulae from the oviduct. Sometimes, if no morulae are flushed
through, it is hard to tell whether the oviduct has been correctly
there is much debris floating around, then the oviduct has been properly
flushed. The morulae are then collected using a mouth pipette and
petri dish in microdrops of M16 medium. These drops are covered
fluid 200 (Dow Corning, viscosity 50CS) and placed in a 37 °C incubator
buffered with 5% CO2.
- Blastocyst transfer is usually performed 24 hours after aggregation when
morulae have become expanded blastocysts, on the same day as the
injection. A little time is given between injection and transfer to
blastocysts to re-expand.
- Careful selection of the recipient is most important, since the pups are
result of a lot of hard work. Two strains of mice are used. RB Swiss
(CBA*C57BL6/J)f1. RB Swiss are quiet and make excellent mothers but
become overweight quickly and do not take anesthesia well. CBA*C57
are hardy and display hybrid vigor. They do not carry excess weight
go under anesthetic well. This strain can be very nervous when
housed separately, which could be dangerous to their young. They are
suitable if a young RB Swiss is placed into the cage as a companion
can be removed as soon as the pups are 7 days old. By this age,
destruction of the litter is very unlikely. If the CBA mother is to be
alone, she must not be disturbed for 10 days.
Prior to surgery, sterilize the following:
- 3 pairs of curved forceps
- 2 pairs of iris scissors
- 1 pair suture clamps
- 1 serafin clip
- Sterile suture with needle attached (small—for mouse surgery)
- Michelle clips (small size)
- Michelle clip applicators
- 1 mouth pipette and flame-polished transfer pipette
- You will also need an anaesthetic. Rompun/Ketavet is found to be quite
effective. To make up 10 mL:
- 0.5 mL 2% Rompun (20 mg/mL Xylazine)
- 0.5 mL 100 mg/mL Ketavet 100—Delta Veterinary Lab, 8 Rosemead Rd Hornsby NSW 2077
- Make up to 10 mL with PBS
- The dosage is 0.02 mL/g body weight.
- Store wrapped in tin foil at 4 °C
- Shake well before use, as it tends to separate in the fridge
DNA Transaction of Eukaryotic Cells Using Calcium Phosphate
- Select a mouse that is 2.5 days pseudopregnant and weigh. Do not use
anything lighter than 25 g or anything heavier than 30 g. Underweight
tend to reabsorb the embryos, as they are not physically ready to
support a pregnancy. Overweight mice make surgery difficult since the
absorption of anesthetic into the fat reduces the potency of the
the presence of fat means the presence of blood vessels, and cutting
through all the extra fat causes a lot of unnecessary bleeding. This
difficult to see what you are doing and may also clog up the tip of your
- Anesthetize the mouse with Rompun/Ketavet, administered
administering the anesthetic, put the mouse back into the box
which it came. The mouse will be more relaxed when placed in a
familiar environment and the anesthetic will act more quickly than it
on a distressed mouse.
check that the mouse is fully anesthetized, press or squeeze the pads
the feet. If the mouse can feel this, it will try to withdraw its leg
grasp. Do not commence surgery until there is no reflex reaction to
- Take the anesthetized mouse and shave its lower back. Lay the mouse on
belly on a petri dish lid, taking care to keep the airway clear by
teeth on the edge of the petri dish. This makes it easier to move the
around without having to actually touch it. Swab the shaven area
hibitane or 70% ethanol.
- Instruments should have been laid out. Use 1 pair of iris scissors and 1
of forceps for cutting the skin—call these “outside” instruments. Use
pair of iris scissors and 2 pair of forceps for working inside the
the “inside” instruments.
the outside forceps and scissors, make a small cut (about 1-cm long)
the dorsal midline of the lower back. Through the shaven, moistened
it is fairly easy to see blood vessels. Try to avoid these vessels when
making the incision (see below).
a pair of outside forceps and inside forceps, pick up the skin and
separate it from the body wall. Cut the body wall as indicated about
long, avoiding blood vessels.
the cut has been made in the right place, the ovarian fat pad is easily
visible. If not, the fat pad can be located by lifting the edge of the
and scouting around with the other pair of inside forceps. Once you
located the fat pad attach the serafin clip to it, taking care not to
ovary. It is important not to damage the ovary, as it is responsible for
hormone production throughout pregnancy. Gently ease the ovary, oviduct,
part of the uterus out through the incision in the body wall. Do not
Pull. A traumatized uterus will contract and move quite violently, making
surgery difficult, and may cause expulsion of the transferred
the tip of the uterus is visible, rest the serafin clip across the
mouse’s back to hold the uterus in place. If the uterus or uterine horn
continually slip back into the cavity, it may be necessary to gently lie
on the side, being careful not to block the airway.
transfer pipette should now be loaded. Five or six embryos must be
transferred to each horn; any less than this and the chances of a
resulting are serverely reduced. It may be loaded in such a way to suit
yourself, but this is a method that is popular. Take up a minute amount
DMEM with Hepes in the very tip of the transfer pipette, then make a
bubble by taking up a little air. Then take up some more medium—
roughly the same volume as you hope to transfer the blastocysts in. Take
another bubble, the same size as before. Then take up your blastocysts
the smallest possible volume of medium, lining them up side by side in
transfer pipette. This is how your transfer pipette should look when
will take some practice. Make sure that you are competent at loading
transfer pipette before any attempt at a blastocyst transfer. During
not the time to learn how to load a pipette. If it is likely to take you
few minutes to load the transfer pipette, then do not expose the
uterus until the pipette has been loaded. This prevents drying out and
further trauma to the uterus. Alternatively, the uterus, ovary, etc.,
moistened repeatedly with a sterile cotton bud and saline.
the pipette is loaded and the uterus positioned, move the petri dish
supporting the mouse to the microscope and turn on the overhead light
source. Once the lights and focus have been adjusted and the mouse
positioned to suit yourself, gently grasp the top of the uterine horn
pair of inside forceps. While still holding the horn with one hand, use
other hand to gently insert a 26-gauge hypodermic needle through the
uterine wall (close to the oviduct) and into the lumen. Choose an area
uterus that is relatively devoid of blood vessels, as blood will clot in
tip of your pipette and block it. Remove the needle carefully (so as not
lose sight of the hole), without averting your eyes, pick up the loaded
transfer pipette. Gently insert the transfer pipette about 3 mm into the
uterine lumen. Gently blow the blastocysts into the uterus, using the
bubbles in your transfer pipette to monitor the transfer. Be careful not
any air into the uterus. Once transfer is complete, quickly rinse out
transfer pipette in some HEPES buffer medium (M2) and check to see if
were any blastocysts stuck in the transfer pipette. If there were,
these blastocysts again.
- With the transfer complete, the serafin clip can now be removed and the
uterus gently eased back into the body. Do not touch the uterus, but
back by the edges of the incision in the body wall and allowing the
fall back in, without actually handling it. This procedure is then
the other uterine horn. The incision in the body wall is not sutured.
is closed with Michelle clips—2 per incision is usually sufficient.
Michelle clips are used on the skin instead of sutures, because the mice
chew at the suture thread and effectively open their wounds up.
- Once surgery is complete, the mouse is placed in a box of clean
sawdust. Under anesthetic, mammals are unable to retain heat as
when conscious. For this reason, and also because the animal has been
shaven, the mouse should be wrapped in a tissue to help keep it warm.
will also be used as bedding until the fur has grown back. It is also
important that the mouse be housed alone, as anesthetized animals
often buried by cagemates who think they are dead. All animals should
recovered sufficiently from the anesthetic before being returned to the
animal room and left unattended. The cage should be placed on a shelf
from male mice, as strange male pheromones will often cause females
abort. Recipient mice should be handled with care and tiptoed around
pregnant mice are easily upset, sometimes leading to abortion, or even
cannibalism of pups.
- M CaCl2 Merck. Filter (to remove particulates), autoclave, and store at
–20°C in aliquots.
- 2X HEBS gm/500 mL
- NaCl 8.0
- KCl 0.38
- Na2HPO4.7H2O 0.19
- Glucose 1.0
- HEPES 5.0
- Adjust pH to 7.05 ± .05 with NaOH, autoclave, and store frozen in
- 1 X HEBS/15% glycerol. 50 mL 2X HEBS)
- 15 mL glycerol
- 35 mL DDW, autoclave and store at –20°C in aliquots.
- Carrier DNA. Mouse liver DNA, sheared at 90 µg/mL in 0.2X SSC by
passing 2X through a 20-gauge needle; filtered through a 4.45-mm filter.
Preparation of CaPO4 Precipitates
- Plate cells at 1X 106 in 10 cm dishes.
- The next day, perform transfection.
Cell Culture from Whole Mice Embryo (Day 10–11 PC)
- Set up 2 rows of tubes—A & B
- In tube A place:15 ug of plasmid DNA (linearized, phenol-extracted,
precipitated, and resuspended at 1 µg/mL in sterile, 0.2X SSC, 69 µL 2M
CaCl2, 460 0.2X SSC.
- In tube B, place 550 µL 2X HEBS.
- Using an autopipette and a 1-mL pipette, have tube B bubbling while
slowly adding contents of tube A.
- Stand 15–20 mins, while precipitate forms, producing a milky fine pipette.
- Carefully add precipitate dropwise to a 10-cm dish of cells while maintaining
the pH of cultures.
- Leave 3–4 hrs in CO2 incubator (Can be left overnight).
- Glycerol Shock:
Aspirate medium: Wash by adding ~3 mL medium then aspirate. Add 2.5
mL 15% glycerol in HEBS.
mins at room temp., then aspirate. Wash with 5 mL medium per
plate. Add fresh medium.
are fragile and only loosely attached at this stage. Handle very gently.
pipette was left on cells overnight, do the splits the next day.
- Trypsinize to harvest cells. Recover cells in medium containing
divide directly in the ratio 3/5 and 1/20th into two 10-cm plates,
- Feed it every 2–3 days with a medium containing antibiotic (once a week
when most cells have been killed).
- Sterile technique
- Autoclave 1 X PBS
- Dissection tools (watchmaker forceps X 2–3, surgical scissors)
- Sterile dishes (10 cm)
- Prepare 6 well dishes to culture the embryo cell culture and mark the well.
- Prepare syringe contain required amount (300 µL) of trypsin and keep in
- Stopcock 3-way transfusion
PCR Genotyping from the Embryo Yoke Sack
- Kill mice.
- Open abdomen and remove the both side of the uterus—sterile from now!
- Open uterus and separate embryo (include decidua).
- Take single embryo into fresh dish, open decidua, open and remove york
for PCR genotype (soak in PBS of the 6 well dishes, wash twice, and
into DNA buffer).
- Rinse the forceps, take the embryo into a fresh syringe, carry the
contain the embryo into a tissue culture hood. Place on a 3-way stopcock
add on a syringe containing 300 µL trypsin on the other side. Start
timing before pushing the syringe. Push the syringe from one side at a
carefully close the stopcock bit by bit (not too much as the cells will
lysed). This requires about 20 seconds (shorter is better), and place
syringe in one well, then place the dish on a warm plate, up to 3
- Rinse the syringe with 1 mL media and add total media of 3–3.5 mL. Do
use a pipette. Take up and push down the culture, since the embryo
tend to be happier when the cells are aggregated.
Incubate the culture at 37°C, 5% CO2 for 3–5 days until the
confluence. Transfer the cell into a 10-cm dish by 1 mL trypsin and 8–10
Prepare the DNA for PCR
- DNA buffer:1 X PCR buffer
- TISSUE CULTURE TECHNIQUES 573
- 0.45% NP-40
- 0.45% Tween-20
- Proteinase K 100 mg/mL
PCR Reaction (50 mL) Contents
- In a PCR tube containing 50 µL DNA buffer.
- Dig washed york sack into tube, spin down.
- Incubate at 50°C for 30 minutes.
- Boil the lysate for 5 minutes, spin down.
- Take 5 µL for PCR.
- 1 X PCR buffer
- 2 mM MgCl2
- 100 nM dNTPs
- 0.25 mM
- 0.05 mM
- 0.2 mM
- a c 400bp (targeting region)
- a b 200bp (wild type)
- 2.5 unite/100 µL Taq DNA polymerase
- Initial denaturing by 94°C for 4 minutes
- 28 cycles of 94°C for 1 minute
- 60°C for 1 minute
- 72°C for 1 minute
- Final extension by 72°C for 10 minutes.