Culture of Postimplantation Mouse Embryos

Paul Martin and David L. Cockroft

1. Introduction

A major disadvantage of working with postimplantation mammalian embryos is their relative inaccessibility to experimentation while they develop within the maternal uterus. Two techniques allow us to get around this problem to a large extent. The first, which is the subject of this chapter, can be used for mouse embryos explanted between 7.5 d of gestation (E7) and 12.5 d of gestation (E12), and involves dissecting embryos from the uterus and culturing them in roller bottles (1). In this way, embryos can be surgically or chemically manipulated or labeled and will develop quite normally in culture for periods of 12-60 h, depending on the stage at explantation (2,3). The second technique, which allows experimentation on more advanced stages, is that of exo utero or open uterus surgery, in which fetuses are suspended in the fluid-filled abdominal cavity of the female mouse while retaining their placental attachment to the uterine wall (4). This procedure is suitable for fetuses at 12.5 d of gestation (E12) and beyond. This chapter will focus on the techniques for culturing E11 mouse embryos with open yolk sacs at limb-bud stages (5) and will also include protocols for culturing earlier stage embryos. We will describe studies in the mouse, but similar manipulations are possible with rat embryos, bearing in mind that they are generally 1 or 2 d behind mouse development; for example, an E11 rat embryo closely resembles a mouse embryo between E9 and E10.

2. Materials

1. Microscope: A good-quality dissecting microscope is essential, preferably with both transmitted (bright- and dark-field) and incident (e.g., fiber-optic) illumination—we use Wild microscopes with an overall magnification range of 6-50x.

2. Instruments: Table 1 shows the tools you will need and what they should be used for.

From: Methods in Molecular Biology, Vol. 97: Molecular Embryology: Methods and Protocols Edited by: P. T. Sharpe and I. Mason © Humana Press Inc., Totowa, NJ

Table 1

Tools for the Job



1 Pair coarse scissors, 1 pair coarse forceps

1 Pair fine scissors

2 Pairs fine serrated, or watchmaker forceps 2 Pairs watchmaker's forceps (no. 5), carefully ground to a tip diameter of 0.05-0.1 mm (see Note 1) Iridectomy scissors (large)

Iridectomy scissors

(small) Hemostat (Spencer-Wells forceps)

Opening the abdominal cavity, when explanting embryos or bleeding rats Removing uterus Removing uterus, opening uterus, exposing aorta during bleeding Removing decidua of E7-E9 embryos, opening Reichert's membrane

Removing decidua/Reichert's membrane of E10-E11 embryos Opening yolk sac of E10-Ell embryos Holding colon out of the way during bleeding

Can also be used for removing uterus

Do not use for opening uterus and so on, since they are too delicate and easily damaged by such heavy work Can use watchmaker's forceps instead

3. Dishes: 5-cm plastic Petri dishes (Sterilin) are suitable for all the operations of explanting embryos at all ages described here. You will need 3-6/litter (more for older embryos); always transfer embryos to a fresh dish if the medium becomes cloudy with blood, and so forth, and ensure that dishes contain enough medium to submerge embryos completely.

4. Saline recipes—see Table 2—we use a 1:1 mix of Tyrode's and Earle's salts (=explanting saline) for explanting open yolk sac embryos. To this, we add extra bicarbonate and glucose (=culture saline) when it is used in the culture medium, in order to equilibrate it with the CO2 in the gas mixture used during culture and to increase the energy sources.

Embryos cultured with closed yolk sacs are explanted in PB1, and cultured in pure rat serum. The pH of all media should be about 7.2.

5. Rat serum: This is the basic culture medium for all postimplantation embryo culture, and is used undiluted for embryos cultured with closed yolk sacs (E7-E10 mouse) (6), or diluted to 25% for culture with open yolk sacs (E10-E11 mouse) (5). Rat serum is available commercially, at a price, but because of the method of preparation, it is inferior, particularly for the culture of the earlier stages (7,8), to what you can prepare yourself (see Subheading 3.1.3.).

Table 2 Media



PB1 medium,

saline, g/L

saline, g/L

































Na pyruvate





Fetal calf serum


10% v/v

6. Sterility: Provided all dishes, instruments, and media used are sterile, we find that explantations and experimental manipulations can be performed on the open bench without problems of infection.

3. Methods 3.1. General

3.1.1. Obtaining Timed Pregnant Animals

The best way to obtain embryos at precisely the stage of development needed is to oversee the mouse mating procedure yourself. If only small numbers of litters are required (1-10 litters/wk), then a colony of 5 studs, with virgin females bought in as needed (say 20 females/2 wk) is quite satisfactory. For larger numbers of litters, it is probably more economical to set up your own breeding colony, with weaned newborns grown up to replace the female stock. We usually mate female mice at 8-12 wk old, and retire our breeding studs before they are a year old. With albino varieties of mice, it is relatively easy to detect the one day in the cycle of 4 or 5 d that females are in estrus, because the vagina appears pink and swollen by comparison with nonestrus females. In nonalbino strains it is harder to distinguish estrus, and if this cannot be judged, then three or four times as many breeding pairs should be set up to make allowance for the nonestrus females. We introduce females individually to studs late in the afternoon, and check for vaginal plugs the following morning. The hard whitish plug occluding the vagina is usually very obvious, but occasionally it is located deep, and this can be checked using a round-ended probe. If you have too few studs for paired matings, then it is possible to put up to three females with one stud, but plugging efficiency and litter number will sometimes be compromised. Generally, we expect 30-80% of our females judged to be in estrus to become pregnant. Depending on strain of mouse, the litters will contain from 5-15 embryos.

3.1.2. Killing Pregnant Animals

In the United Kingdom there are "Home Office" (HO) regulations stipulating methods of killing animals. The most satisfactory HO Schedule 1 method for killing a pregnant mouse is by cervical dislocation. This is done humanely by placing the fingers of one hand behind the animal's neck and stretching it by pulling the mouse's tail with your other hand. Alternative methods of killing include overdoses of anesthetic, but this may have adverse effects on the later health of the embryos.

3.1.3. Making Rat Serum

Rats (preferably fasted overnight) should be anesthetized with halothane or (if permitted) ether. Lay the anesthetized animal on its back, douse it with 70% ethanol, and open the abdominal cavity. Displace the small intestines to the left, pull the colon to the right, and hold it there with a hemostat (Spencer-Wells forceps). Expose the dorsal aorta with fine forceps (it is generally smaller and paler than the adjacent vein), and insert, bevel down, a 19- or 21-gage needle connected to a 10- or 20-mL syringe, with all air excluded. Gently withdraw blood until the rat stops breathing, after which only a further milliliter or two can be obtained. Remove the needle and transfer the blood to a 12- or 15-mL centrifuge tube, running it gently down the side of the tube. Immediately spin the tube for at least 5 min at 2000g. This will precipitate the blood cells, and a whitish clot will form in the supernatant serum. When all rats have been bled (20-30 is a reasonable number for a single session), squeeze the clots with long slender forceps to expel serum, spin again as above, and decant the serum into 50-ml centrifuge tubes with a pipet. Spin again to bring down any persisting blood cells, and then pool the day's serum in a suitable container (e.g., a 250-mL tissue-culture flask). Add antibiotics if desired (e.g., 100 |g/mL Streptomycin, 100 IU/mL penicillin), and aliquot the serum in quantities of 5-20 mL before freezing. It will keep for several months at -20°C, or years at -70°C (freeze first at -20°C, and then transfer to -70°C; otherwise tubes may crack). See refs. 3 and 9 for further information on rat bleeding.

3.1.4. Preparing Rat Serum for Culture

Thaw the serum at room temperature or in a 37°C water bath, then heat-inactivate it for 30 min at 56°C. Gassing the serum while hot will help drive off any persisting anesthetic dissolved in it. A residual clot often appears at this stage, and can be removed by centrifugation or, more effectively, by filtration through a 0.45-^m syringe filter (Sigma). Serum is then ready for use, either pure or diluted, according to embryonic age.

3.2. Explanting E11 Embryos (Cultured with Opened Yolk Sacs)

1. Although the procedure for explanting with open yolk sacs described and illustrated here is for E11 embryos, it is equally applicable to E10 embryos. The freshly killed pregnant animal is laid on its back and the abdomen doused with 70% alcohol. The abdominal skin can be pinched between thumb and forefinger of both hands, and the skin torn back towards head and tail to reveal a clean peritoneal surface, which can then be opened with scissors and forceps. This method avoids contaminating the abdominal cavity with hair. Alternatively, all abdominal layers can be opened at once with a large U-shaped scissor incision, with the two ends of the U at the hindlimbs.

2. The uterus with embryos is then lifted clear of the abdominal cavity. It can be held with forceps midway along one horn at a site between two embryos and needs to be severed with scissors at the tip of that horn (near the ovary), then where it communicates with the cervix (without separating the two horns), and finally at the tip of the second horn. As this is being done, the uterus can also be trimmed free of mesentery and fat.

3. The whole uterus should then be rinsed in PBS and remaining mesentery cleared away before transfer to a fresh dish of PBS. The uterus can now be opened to expose the embryos. This is done by carefully tearing along the antimesometrial wall of the uterus with fine forceps (Fig. 1). It is easy to damage embryos at this stage, and the best way to avoid this is to keep the two pairs of forceps close to one another, so that as they tug apart, all the effort goes into tearing the uterus and not squashing the embryos. Once the uterus is open, the embryos (concep-tuses) appear like peas in a pod attached to the uterus only in their placental regions. The easiest way to free this attachment is to hold the uterus tightly with one pair of forceps, slip the other pair of forceps on either side of the uterine sheet, and then drag them across each placental attachment in turn, gently teasing the embryos free (Fig. 2).

4. The individual conceptuses can now be transferred by pipet to a fresh dish of explanting saline, and their decidua removed. The decidua of an E11 embryo is thin and is best peeled off, beginning on the side opposite the placenta, by shallow pinching with two pairs of fine forceps, which allows a gentle tearing action. Often the very thin and transparent Reichert's membrane, which sits beneath the decidua and clings to the yolk sac, will rupture during this operation, in which case it is easy to remove it with the decidua. When the decidua has been torn back to about the level of the placenta, it can be tidied up by trimming off with a pair of iridectomy scissors (Fig. 3). If Reichert's membrane is still intact, it must be torn open and trimmed back to the placenta also.

Fig. 1. The uterus (Ell) is gently torn open along its antimesometrial wall using fine forceps to expose the conceptuses.

5. Next, the yolk sac must be cut with fine iridectomy scissors, close to where it abuts the placenta, taking care to avoid damaging any of the larger yolk sac vessels (Fig. 4). First use two pairs of watchmaker's forceps to make a small hole in the yolk sac adjacent to the placenta, to allow access for the iridectomy scissors. The yolk sac should not be cut completely free of the placenta—rather cut about 4/5 of the way around, creating enough of an opening for the embryo to be pulled out of the yolk sac head first. The region

Fig. 3. The decidua and thin Reichert's membrane overlying the E11 yolk sac are trimmed back level with the placenta.

Fig. 4. A small hole is made in the yolk sac close to where it abuts the placenta and adjacent to the forelimb bud. Iridectomy scissors are then used to cut around 4/5 of the base of the yolk sac.

of yolk sac left uncut should be that adjacent to the tail of the embryo— otherwise delivery is made significantly more difficult. The embryo is drawn out of the yolk sac by pulling on the amniotic membrane overlying the embryo's head with one pair of forceps, while holding the mouth of the yolk-sac incision with a second pair of forceps (Fig. 5). After the head is outside the yolk sac, it is necessary to rupture the amniotic membrane in order to exteriorize the rest of the embryo. Last of all, the yolk sac and amniotic membranes are flipped under the tail, and the embryo is now available for manipulations or immediate culture.

Fig. 4. A small hole is made in the yolk sac close to where it abuts the placenta and adjacent to the forelimb bud. Iridectomy scissors are then used to cut around 4/5 of the base of the yolk sac.

Fig. 5. The E11 embryo is delivered from the yolk sac by tugging on the amniotic membrane overlying the embryo's head.

6. If the developmental stage of the embryos is critical for your experiment, you should stage them now (see Note 2).

7. All of the above procedures and any planned subsequent manipulations (see Note 5) should take no more than about 2 h, or subsequent development in culture will be compromised. This generally imposes a limit of no more than 2 litters of embryos/culture session. When embryos are ready for culture, they are individually transferred into 50-mL Falcon tubes containing 5 mL of 25% (v/v) rat serum in culture saline (see Table 2 for recipes). To ensure a gaseous seal, we apply a thin coat of silicone vacuum grease (Dow-Corning) to the rim of the tube. The tube is then gassed with 95% O2, 5% CO2 (for 1 min at a gas flow rate sufficient to ruffle the surface of the medium gently—Fig. 6). The Falcon lid is then screwed tight and the tube is placed in a 37°C incubator containing rollers-rotating at 30 rpm. We find that the custom-made BTC roller-incubators (see Note 9) are excellent for this purpose. By carefully stacking Falcon tubes in the incubator, it is possible to culture up to 18 tubes (embryos) at one time.

8. The health of embryos can be determined at any stage during culture by brief removal of the Falcon tube from the incubator and viewing under a dissecting microscope with transmitted light. Because the walls of the tube are translucent, it is easy to check how well the heart is beating, and usually whether there is good circulation in the larger vessels of the yolk sac and over the brain. Also, check whether there is any swelling of the pericardium or blistering of distal limb ectoderm (usually signs that the embryo is faring badly). In any case, embryos should be regassed every 12 h or so. Only badly damaged embryos will fail to culture successfully for 12 h and, of these, 90% will successfully make it through to 24 h: From this stage on, survival rates get steadily worse with only 1 in 3 embryos

Fig. 6. Embryos are transferred to a roller bottle containing culture medium and gassed with 95% oxygen before culture begins.

healthy at 36 h and 1 in 6 or 7 making it through to 48 h. Figure 7A shows the appearance of E11 embryos before and after culture for 24 h.

Explanting E9 Embryos (Cultured with Closed Yolk Sacs)

Embryos younger than E10 at explantation are cultured with the visceral yolk sac intact. The initial stages of killing the mouse, opening the uterus, and separating the conceptuses are the same as for the E11 embryos, though of course the con-ceptuses are smaller.

Removal of the decidua of E9 embryos is similar to the E11 procedure, since it already forms a relatively thin layer over the conceptus. It is advisable to remove all of the decidua, starting at the equator and tearing toward and over the placenta, but being careful not to damage the latter, since it has an extensive blood circulation at this stage.

Next, open Reichert's membrane; although this is thin and transparent, it is mostly overlain with a layer of trophoblast and blood cells, and it may be necessary to pick through this layer before rupturing Reichert's membrane, after which removal is straightforward, trimming it up to, but not beyond, the placental border (Fig. 8).

Fig. 7. (A) Two mouse embryos taken from the same uterus at E11. The one on the left was refrigerated at explantation. The one on the right was cultured with open yolk sac as described for 24 h before photography. (B) E9 embryos before and after culture.
Fig. 8. The Reichert's membrane of the E9 embryo is opened using watchmaker's forceps, and the surplus is trimmed to the border of the placenta.

4. If the visceral yolk sac or placenta is damaged, causing deflation or bleeding, the embryo should be discarded; otherwise, it is ready for culture.

5. E9 embryos may be cultured in 50-mL Falcon tubes as above or 30-mL Universal containers (Nunc, with a smear of silicone grease round the internal rim of the lid to provide a gas-tight seal) with 1-1.5 mL pure serum/embryo, and 2-5 embryos/ tube, depending on size. Initially, the embryos should be gassed with 5% CO2 in air (i.e., 20% O2, 75% N2, 5% CO2), which is replaced with 40% O2, 55% N2, 5% CO2 after 18-24 h of culture. Figure 7B shows the extent of development of E9 embryos after 48 h in culture, though 18-24 h are a more realistic period if you need the majority of embryos to be healthy at termination.

3.4. Explanting E7 Embryos (Cultured with Closed Yolk Sacs)

1. The initial stages are as above, but with E7 embryos, removal of the decidua requires a rather different strategy, since the conceptus is embedded in a relatively thick mass of decidua. Start by impaling the decidual mass parallel with its long axis, but off-center, with one arm of a sharpened slender pair of forceps. Then squeeze the forceps together to form an incision in the decidua.

2. Repeat on the other side of the decidua, then unite the two incisions around the base of the conceptus (usually the thicker end).

3. Now grasp the two flaps of decidua with two pairs of watchmaker forceps, and pull apart (Fig. 9), when the conceptus should be exposed, and usually it will

Fig. 7. (continued) The embryo on the left was refrigerated at explantation; on the right is a littermate cultured for 48 h. The embryos are shown dissected free of the extraembryonic membranes with which they are cultured. (C) E7 embryos before and after culture. An E7 conceptus (including the visceral yolk sac and ectoplacental cone with, which it would be cultured) is shown on the left. This was refrigerated while littermates (center—with membranes as cultured; right—dissected free of membranes) were cultured for 48 h. Scale bars for A, B, and C are all 1 mm.

Fig. 9. As the decidua of the E7 conceptus is pulled apart, the ectoplacental cone of the embryo within can be seen.

remain on one of the decidual halves as they are separated. If the embryo sticks to both decidual halves, so that separating them further might damage it, either hold the flaps apart with one pair of forceps, whilst teasing the embryo free on one side with the apposed tips of the other pair of forceps, or make a further incision in one of the decidual halves, so that only a quarter is removed initially, followed by the second quarter.

4. Once this has been accomplished, further divide the base of the decidua containing the embryo along the long axis, and peel apart along the length of the conceptus (Fig. 10). Repeat if necessary until the embryo is attached only to a thin sliver of decidua, like a segment of an orange.

5. Impale the sliver of decidua on either side of the embryo with the points of one pair of forceps, stretching it slightly, and tease the embryo free with the apposed tips of the other pair of forceps (Fig. 11).

6. All that remains is to open Reichert's membrane (Fig. 12); as with the E9 embryos, this may be overlaid with blood and trophoblast, though the membrane itself is thin and transparent. Sometimes Reichert's membrane stands clear over the embryo (at the end opposite the reddish ectoplacental cone), where it can be grasped and pulled apart. Otherwise, it will be necessary to use two pairs of watchmaker forceps to rupture the membrane midway along the length of the conceptus, in a region overlying the visceral yolk sac, where unseen damage to the underlying tissue will be less serious than damage to the embryo itself. Once opened, Reichert's membrane is trimmed off to the border of the ectoplacental cone, which is left intact.

7. The E7 embryo is now ready for culture. These embryos are cultured with 1 mL pure serum/embryo, with three embryos per 30-mL Nunc tube, or up to five embryos per 50-mL Falcon tube. They are gassed initially with 5% O2, 90% N2, 5% CO2, and then with 5% CO2 in air (i.e., 20% O2) after 24-36 h of culture (i.e.,

Fig. 10. The decidua is removed by successively peeling off strips along the long axis of the conceptus.

Fig. 11. The conceptus is teased free from a narrow sliver of decidua.

Fig. 11. The conceptus is teased free from a narrow sliver of decidua.

Reichert's membrane is opened with finely ground watchmaker's forceps.

when the heart beat and visceral yolk sac circulation are established). Figure 7C shows the extent of development of E7 embryos after 48 h in culture.

4. Notes

1. Sharpening forceps: A combination Japanese waterstone (1000/6000 grit, e.g., King brand) is ideal. Wet it with distilled water and grind the forceps along their length until a tip diameter of 0.05-0.1 mm is achieved. It is very important that the tips meet precisely, without crossing or one protruding beyond the other. Then round off all but the innermost (mating) edges, so that when apposed, the tips together form a blunt-ended probe.

2. Within and between litters of embryos of the same age, the range of developmental stages can vary dramatically. There are a number of established staging guides: ref. 10 covers all stages from fertilization to birth, ref. 11 from fertilization to 4 wk postpartum, ref. 12 from 8-16 d, ref. 13 from 7.5-10.5 d, ref. 14 from 6.75-8.0 d, and ref. 15 from 9 d to post natal stages. Between E9 and E14 a useful method of accurately staging embryos is to use the shape of the fore- and hindlimbs (16). These publications will also be useful in assessing the stage of development reached after culture and how it compares with growth in vivo. References 10 and 12 also provide a wealth of additional information, including sectioned embryos, which can help determine the success of your experiment.

3. An alternative method of transferring embryos from dish to dish while they are surrounded by their decidua is to pick them up with watchmaker forceps where the decidua is thickest, thus minimizing carryover of medium from one dish to another.

4. When the time comes to harvest your cultured embryos, they should be gently slipped into ice-cold PBS, and trimmed free of their yolk sac and placenta by cutting the umbilical vessels. Embryos can then be transferred to the fixative of your choice. We favor half-strength Karnovsky's fixative (17) if embryos are to be subsequently processed for scanning electron microscopy or resin histology, or 4% paraformaldehyde if they are for whole-mount or sectioned immunocy-tochemistry or in situ hybridization studies. For general wax histology, good results are obtained with Bouin's fixative followed by staining with Hematoxylin and Eosin after sectioning (10).

5. Manipulations: We have successfully performed a number of manipulations on the E11 embryo after it has been delivered from its yolk sac and prior to culture. It is possible to inject reagents into various epithelial lumens—for example, we have injected the marker dye Monastral Blue into the otic vesicle (18). Similarly, it is possible to perform simple surgical operations (19-21) and to label groups of cells with the lipophilic dye, DiI (21). Even if the embryos bleed a little after surgical manipulations, they generally survive and can be successfully cultured. The two biggest problems with any such manipulations are being able to see well enough what you are doing and keeping the embryo from moving around the dish as you operate on it. The first of these problems is easily resolved with good lighting (both incident and transmitted) and the second requires gentle holding or supporting of the embryo with forceps.

6. Transgenic mice: It is possible to culture transgenic mice, but these will generally be derived from heterozygote crosses, so only a fraction of the embryos cultured from each litter will be of the required genotype. It is sometimes possible to take a small piece of tissue (tail tip or a small piece of yolk sac) at the time of explant, so that embryos can be genotyped by PCR during the culture period. Otherwise, this can be done with spare embryonic tissue taken after the culture period, but before fixation.

7. Synthetic medium: We have recently begun to culture E11 embryos with open yolk sacs in a serum-free medium made by Gibco BRL. This medium is excellent for E11 cultures up to 24 h, but we have not tried it for longer culture periods, or its efficacy for the more sensitive younger embryo stages. Good results can also be obtained over a range of stages with partially defined medium, made by extensively dialyzing rat serum, and then supplementing it with glucose, vitamins, and amino acids (22).

8. Adding glucose to serum: If you have fasted your rats before bleeding, they will have lowered blood glucose levels. Addition of 8 |L/mL of a 50 mg/mL stock solution of glucose to the serum will restore the glucose levels to that in serum obtained from fed rats.

9. Address for BTC: BTC Engineering, 12 Shirley Close, Milton, Cambridge CB4 4BG, UK.


We are particularly indebted to Catherine Haddon for the lucid diagrams illustrating the dissections. We also thank The Medical Research Council and the Wellcome Trust (P. M.), and the Imperial Cancer Research Fund (D. L. C.) for financial support.


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