Timed matings to produce host embryos and pseudo-pregnant foster
mothers
1. Day 1: Embryo donors are intraperitoneally (IP) injected with 5 IU of PMSG
using a 1 mL syringe with a 26G needle. PMSG is diluted in 0.9% NaCl. In one
experiment, ten embryo donors and ten stud males are used.
2. Day 3: 42–48 hr after administration of PMSG, the same females are IP injected
with 5 IU of hCG and mated to singly housed males of proven fertility.
3. Day 4:
a. Copulation plugs are checked the in the morning. The copulation plug is a
coagulation of seminal fluid and protein and is visible in the vagina of the
female. Place all of the females into a cage and mark the day the animals were
plugged. This day is considered 0.5 days post-coitum (E0.5).
b. Females are checked for natural oestrus and are mated to singly housed
vasectomized males. In order to ensure 10 pseudo-pregnant females, it will
be necessary to examine approximately 100 animals for oestrus. An oestrus
female may be identified by the degree of swelling, pink color and moistness
in the vaginal area. Vasectomized males can be conveniently purchased or
generated by surgical procedures described elsewhere (Bradley, 1987).
4. Day 5:
a. Check for the presence of a copulation plug in the pairings with vasectomized
males.
b. Place the females who have successfully mated, i.e. a copulation plug is clearly
visible in a separate cage. Indicate on the cage card the date of the plug.
B. Collection and culture of blastocysts and pre-compacting morulae
Collection of E2.5 eight-cell stage host embryos
1. Set up microdrop cultures as follows:
a. Microdrop cultures are set up by arranging several 20 L drops of G2v3 culture
medium in 60 mm dishes and quickly but gently overlaying with embryo
culture grade mineral oil.
b. The dishes are equilibrated in a tissue culture incubator at 5% CO2 for at least
30 min before use.
2. Humanely sacrifice a superovulated embryo donor mouse in the morning of E2 and
lay it on its back. Rinse with 70% ethanol. Pinch the skin with micro-dissecting
curved forceps and make a lateral incision in the midline area of the abdomen
using fine scissors.
3. Holding firmly above and below the incision, pull firmly in opposite directions
until the abdominal area is fully exposed.
4. To expose the reproductive tract, cut open the peritoneum to reveal the contents
of the abdominal cavity. Push the intestines to one side to reveal the U-shaped
uterine horns with the ovary and oviduct at the top of each horn.
5. Cut away as a single unit the fat pad (above the ovary and attached to the kidney),
ovary, and oviduct and just below the horn of the uterus and place into M2 medium.
6. In 3 mL of M2 and 2 mL of PBST (0.05% Tween-20 in PBS) cut away the fat pad,
ovary and most of the oviduct using micro-dissecting scissors and micro-dissecting
forceps.
7. Using a pair of micro-dissecting forceps, hold the uterus with one of the pair and
extrude the content therein in the direction of the oviduct with a second of the
pair.
8. Collect and move the pre-compacted eight-cell stage embryos into microdrops
using aspirator tube assembly and a pulled glass capillary pipette. Pulled Pasteur
pipettes are used to move embryos between microdrops and are made by heating
and pulling a calibrated glass capillary pipette over a flame of a micro-burner (see
protocol for preparing glass tools).
9. The embryos can be used at this point for preparing aggregations with human ES
cells or cultured overnight for use in blastocyst injections.
Procedure for collection of E3.5 early blastocyst host embryos
1. Set up microdrop cultures of G2v3 medium as in the procedure for collection of
E2.5 embryos.
2. To expose the reproductive tract, follow steps 2 through 4 in the procedure for
collection of E2.5 embryos above.
3. Cut the horn of the uterus just above the cervix and above the fat pad above the
ovary. Release the uterus by cutting through the mesenteries and place on a piece
of absorbent paper towel. Grasp the uterus and cut away the mesenteries and blood
vessels.
PROTOCOLS 131
4. Move to a dish with M2 medium.
5. Insert a 25G needle and syringe loaded with 3 mL of M2 and 2 mL of PBST into
the oviduct end of the uterus and hold the uterus on the needle with the curved
micro-dissecting forceps.
6. Flush the blastocysts through the uterus towards the cervix end with about 1 mL
of media.
7. Collect and move the early blastocyst stage embryos into microdrops of G2v3
medium using aspirator tube assembly and a pulled glass capillary pipette.
8. The embryos can be incubated at 5% CO2 until used for microinjection of human
ES cells.
Preparation of human ES cells for blastocyst injection
Cells that have been maintained on MEF feeders or on Matrigel in MEF conditioned
medium have been used for these protocols. We have included protocols for the passaging
and harvesting methods that we used to produce cells for blastocyst injections
and morula aggregations. Generally, cells grown on MEF feeders have been used
for blastocyst injections and cells grown on Matrigel have been used for the morula
aggregations. However, we have had success generating chimeras with both passaging
schemes in the protocols. We have successfully used these protocols with human ES
lines derived in house i.e. RUES1 and RUES2.
1. Prepare microdrop cultures as for embryos except use 50 L drops HUESM supplemented
with 20U/mL of DNAse I.
2. Prepare MEF feeder plates if passaging human ES cells. Prior to passaging the
cells by microdissection, medium is changed in the well to be passaged and on a
fresh feeder plate. Protocols for generating and mitotically inactivating feeders are
described in Chapter 4. We have also used a commercial source of Mitomycin-C
inactivated MEFS.
3. Prepare media: We have used both H1 medium and HESM successfully (see
Table 1). To prepare complete growth medium, bFGF is added just before feeding.
To prepare complete growth medium, bFGF is added just before feeding (see
table below). Stocks of bFGF are made to 100 g/mL in sterile 10 mM Tris-HCl
pH7.6 with 0.1% BSA. Aliquots of a convenient volume are frozen at −20C. For
maintenance on MEF feeder layers, bFGF is supplemented to 40 ng/mL before
feeding the human ES cells.
4. Prepare glass tools for microdissection: For glass tools, Pasteur pipettes are pulled
into hair-thin hooks. The hooked end of the glass tool should be thin enough for the
microdissection of the human ES cell colonies (such dissection is also described
at length in Chapter 5) but thick enough to withstand some pressure during the
dissection. Fine glass needles with hooked ends are forged in two steps over a
microburner that can produce a very small flame.
a. Establish a small candle-size flame with a micro-burner made from a Bunsen
burner fitted with an 18G needle.
b. For glass hooks, hold a Pasteur pipette at both ends and melt the glass approximately
one inch below the taper until the lumen is fused and the glass glows
orange. (For pulled pipettes used in embryo manipulations, melt the glass, but
do not fuse the lumen.)
c. Quickly remove the glass from the flame while simultaneously pulling the ends
away from each other. The glass should be drawn out to a filament without
breaking. (For pulled pipettes used in embryo manipulations, stop here and
break the filament with an inner diameter slightly larger than the embryos.)
d. Again, while holding a Pasteur pipette at both ends several inches above the
flame, slowly lower the drawn filament approximately one inch from the new
taper towards the flame while gently pulling at each end. Before reaching the
flame, the filament should melt in two while being drawn into a very fine
filament.
e. While holding the large end at approximately a 90 angle to and several
inches above the flame, lower the filament tip towards the flame. The rising
heat should curl the filament tip up forming a “hook”.
f. Using a no. 5 watchmakers forceps, trim the end of the filament to finish the
end into a clean “hook”.
5. Preparing human ES cells for injection or passaging: Ideal colonies or parts of
colonies are micro-dissected into clumps of cells using the glass hooks. The hook
is used to gently pull apart pieces of the colony. This can also be accomplished
by cutting a grid into the colony with the back of the hook and pulling the pieces
away from the colony one at a time. For passaging, a chunk size of 100–200 cells
is optimal. For injection, move the clumps to a fresh microdrop of media. The
clumps are further dissected into clumps of 10–15 cells using glass needles (see
Figure 2F). The back edge of the glass hook can be used to cut larger chunks into
smaller pieces. Movies demonstrating this technique can be found on the Brivanlou
lab web site (
http://xenopus.rockefeller.edu).
6. For passaging, transfer clumps to fresh feeders: For routine passaging after
microdissection, the cell clumps are swirled into the center of the dish and 20
to 50 clumps are transferred to the new feeder wells using P1000 micropipettes.
Pre-coat the micropipette tip with the medium so that the cells do not stick.
If possible, leave the dishes untouched on a warmed surface (preferably under
O2/CO2 blood-gas mix) for 15–30 min to allow the clumps to begin attaching to
the dish before moving to an incubator. Excessive handling of the new dish will
cause the clumps to migrate to the center of the dish rather than remaining evenly
distributed across the dish. Complete growth medium is exchanged on the growing
PROTOCOLS 133
colonies every day. The lines should culture for no more than 6 days to a week.
The timing of passage is dependent upon the appearance of differentiation within
the colonies — mainly from the center of the colony.
Enzymatic passaging and harvesting of human ES cells for morula
aggregation
Human embryonic stem cells can also be grown in feeder-conditioned medium on a
substrate of extracellular matrix (ECM).We routinely use Matrigel as a growth substrate
for growing RUES1 human ES cells in preparation for aggregation experiments. Here
we describe the methods used to prepare these human ES cells for aggregation and
the passaging conditions used to maintain them. This protocol can also be used to
prepare human ES cells for blastocyst injections. We caution that alternative enzymatic
passaging protocols may not yield successful embryo aggregates.
Before passaging, examine the colonies under the microscope and look for any
colonies that are differentiated. Spontaneously differentiating areas of the culture can
be removed with a glass tool as described in the manual dissection protocol or aspirated
using a pipette attached to a vacuum. Several types of differentiation can be
morphologically identified in spontaneously differentiating cultures. Look for the center
of colonies that show a depression or “crater” appearance. Areas of colonies
that have begun forming cystic structures in the center of the colony should also
be removed. Also avoid the edges of colonies that do not have a tight border between
the feeder layer and the colony. Areas where the human ES cells have started to
flatten and polarize can also be removed. However, some differentiation on the borders
of the colonies can be tolerated, as these cells will detach during the washing
steps. Ideal colonies comprise small, round cells with a high nuclear to cytoplasmic
ratio that are randomly organized and have not begun forming structures within the
colony.
1. Prepare Matrigel coated plates for passaging and maintenance of cell lines.
Matrigel-coated plates can be prepared in a slightly different manner to that
described in Chapter 5:
Matrigel is diluted to 0.333 mg/mL in cold XVIVO-10 medium (Table 2). All plastics
that come into contact with the undiluted Matrigel should be kept as cold as
possible. We pre-cool six-well plates, dishes and P1000 filter-tips at — 20C for about
20 min. Keep the plates on the ice cold platform at all times. Place the entire ice
bucket with plates into the refrigerator to coat overnight. Before plating cells, check
the coating on the microscope for a meshwork-like single layer matrix. When ready to
plate human ES cells, aspirate the Matrigel from the wells using a Pasteur pipette in
the corner of the well. Get as much Matrigel off of the dish as possible leaving a thin
coating on the surface of the dish. Do not scrape the bottom of the dish. Rinsing the
Matrigel-coated plate is not necessary.
2. Prepare conditioned medium for passaging and maintenance of human ES cells:
MEFs for feeder plates can be prepared and inactivated as described in
We have also successfully used a commercial source of Mitomycin-C inactivated
MEFS. Approximately 5 million cells from frozen vials or 3 million cells from
freshly irradiated MEFs are plated on gelatin coated 10 cm plates and allowed
to attach overnight in FM10 media (see Table 1). HESM growth medium is
exchanged and conditioned for 24 hr before use. For maintenance on Matrigel,
bFGF is supplemented to 20 ng/mL before conditioning on MEFs. The feeder
conditioned medium (CM) can be used immediately, stored at 4C for a week or
frozen at −80C. The conditioned medium is further supplemented with 40 ng/mL
before feeding human ES on Matrigel. Passaged human ES cells are plated in 2 mL
of CM per well of a six-well plate. Feeder plates can be used for up to 14 days
to generate conditioned medium.
3. Passaging human ES cells:
a. Replace the growth medium with Dispase or Collagenase (type V) at 1 mg/mL
dissolved in growth medium and sterile filtered.
b. Incubate for about 4–5 min in a tissue culture incubator. Check the progress
of the matrix digestion, beginning at about 4 min. The colony borders will
begin to peel away from the plate, while the center will remain attached (see
Figure 2A, B).
c. Ideally, gently wash the dispase (or collagenase) off of the plate with growth
medium twice. The colonies should remain attached to the plate. If they have
detached after the dispase incubation transfer all of the colonies and Dispase
solution to a conical tube and centrifuge at 150 g (∼1,000 rpm) for 4 min and
wash the colonies with growth medium. They should get two to three washes
total — either on the plate or with centrifugation.
d. If the colonies remained attached after washing, harvest the colonies with a
cell lifter (see Figure 2C, D).
e. Transfer all of the colonies and growth medium to a conical tube and spin to
pellet the colonies at 150 g/∼500 rpm for 4 min.
f. Using the CM, resuspend the colonies using a P1000 pipette tip in about
500–700 L of medium.
g. Triturate the colonies to clumps with an average size of about 100 cells using
the P1000 tip (see Figure 2E).
h. Plate a portion of the clumps at a 1:10 split ratio. However, the ratio will
need to be optimized for the confluence of the starting population. If possible,
leave the dishes untouched on a warmed surface (preferably under O2/CO2
blood–gas mix) for 10 min to allow the clumps to begin attaching to the
dish before moving to an incubator. Excessive handling of the new dish will
cause the clumps to migrate to the center of the dish rather than remaining
evenly distributed across the dish. Good spacing between the colonies will
allow proper growth of the colonies
