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This unit lists and describes protocols used in the production of chimeric mice leading to the generation of gene knockout mice. These protocols include the collection of blastocyst embryos, ES cell injection, and uterine transfer of injected blastocysts. Support protocols in the superovulation of blastocyst donor mice, generation of pseudopregnant recipients, fabrication of glass pipettes, and generation of germline mice are also included. Practical tips and solutions are mentioned to help troubleshoot problems that may occur.
The generation of knockout mice through the process of creating chimeric mice via embryonic stem (ES) cell injection is a powerful tool for understanding in vivo functions of a particular gene of interest. Knockout mouse models may be used to elucidate the molecular mechanisms underlying human diseases and could aid in the creation of new therapies to treat these diseases.
ES cell injection is the most common method used to generate these chimeras, but morula aggregation is also used. There is a very narrow time window in embryonic development for the ES cell injection procedure to be successful in the generation of chimeric mice. For this reason, all components of the procedure for generating chimeric mice—harvesting, injection, and surgical transfer of the mouse blastocysts—are usually done the same day and require good planning and coordination.
Learning all of the procedures involved requires time and practice. It also requires skills not everyone might possess. How fast and easily these techniques are learned will depend on previous experience, natural abilities, and the desire to learn them.
Note: All animal work performed must be approved by the Animal Care and Use Committee regulations of the institution where the work will be taking place. The animal facility where the mice are held should ideally be pathogen free.
Embryonic day (E) 3.5 blastocysts can be found free-floating in the uterine horn. To collect these embryos, the uterine horns are generally flushed out with media. Although this is considered a straightforward technique, it does require a certain amount of manual dexterity to perform this procedure. Another important component is the ability to use a mouth pipette in the collection and washing of embryos. Without this ability it would be hard, if not impossible, to collect the mouse embryos. Even though in theory only blastocysts should be obtained from this procedure, it is not uncommon to obtain earlier stage embryos as well. Of these earlier stage embryos, morulas should be kept in culture along with blastocysts. Depending on the stage of development, the morula may become blastocysts in culture and can be used for ES cell injections if needed. When flushing out the blastocysts, a zoom setting of 1.2 is suggested, since it will allow a viewing of the uterine horn all the way to the opening of the cervix. For washing blastocysts, a zoom setting of 3.2 is recommended in that it offers enough magnification to discern the blastocyst well, but still allows for a relatively large viewing area to see the embryos on the dish.
Note: It is suggested that all 10 blastocyst donor females be used even if a plug is not seen in some of them. These females could potentially be plugged even though a copulation plug was not seen. Failure to use all of the mice could potentially decrease the embryo yield.
Note: If the needle does not move freely, and the uterine wall expands, the needle is not inside the lumen. A good sign that the uterine horn is being flushed out well is that the medium can be seen coming out of the cervix and the uterine horn expands, showing the rib-like structures of the uterine wall.
Note: When flushing out through the cervix, orient the cervix opening so that it is to the side and slightly up. This allows one to see the media being flushed out and help prevent media from squirting out of the dish. Transfer pipettes used for washing the embryos should be a little larger to allow for capillary action but not so big that there is no control in the mouth pipetting of embryos.
When injecting blastocysts, the optimal type of blastocyst to inject would be between the middle and late stages, with a good-sized blastocoel cavity. Larger, more developed blastocysts tend to reexpand faster after injection but are more susceptible to collapse prior to injection. Trying to inject any hatched blastocysts can be difficult because they lack rigidity from the loss of the zona and have a very sticky outer surface, so they should be avoided. See Figure 3. The morphology of ES cells to inject should be round and smooth and appear opaque in nature. It is not uncommon to have small, medium, and large cell sizes. Medium-sized cells would be the preferred choice in that they will likely give a higher chance for germline transmission, and they are less likely to be differentiated. Avoid the largest cells because they are mouse embryonic fibroblasts (MEFs). See Figure 2. Morula injections could be used instead of blastocyst injections to increase the success of generating chimeras and have higher germline efficiency. The authors have found this not to be necessary. A circulating pump may be used during the cooling stage in place of the dish to cool the ES cells and blastocysts. A suggested microinjection setup is listed in the Materials section. See Figure 9.
The microinjection system should be functioning properly before proceeding with injection of ES cells into blastocysts. Address all problems before starting the actual microinjection procedure. Both injection and holding systems should be in working order before attempting ES cell injections. See Critical Parameters and Troubleshooting for details on problems encountered when injecting blastocysts.
For a cooling system, we use a glass injection chamber with an internal hollow opening around the injection chamber to allow for chilled (4 °C ) water to circulate. See Figure 5. There are 2 reasons to use some sort of cooling apparatus. The first is to delay the reformation of surface proteins after trypsinizing the cells so they do not become sticky. Secondly, cooling aids in the rigidity of the blastocysts. The injection dish should be free of alcohol as this can cause differentiation of the ES cells.
Note: The authors recommend a cell suspension of 1 × 106. This is more than enough cells for injection purposes. If too many cells are added to the injection dish, the cells tend to become sticky. A high ratio of ES cells to feeder cells is best for injections. Having some feeder cells can be a benefit in that they can be used to clear debris from inside the injection pipette.
Blastocysts are more tolerant to pH and temperature changes than that of other earlier stage embryos such as zygotes. Cooling the temperature will delay the onset of blastocyst hatching. Keeping blastocysts in the cold injection chamber too long might be detrimental to their vitality.
When first learning ES cell injections, 200× magnification should be used. Focus on the outer ring of trophoblast cells and then the tip of the injection pipette before attempting to inject inside the blastocoel cavity. This ensures that the injection pipette is in the correct plane of the z-axis, being approximately in the center of the blastocoel cavity. After becoming proficient in ES cell injections, a tapping technique, which looks at indentations on the blastocyst, can be employed to make sure the injection pipette is in the right plane. 100× will work well with this procedure and has the advantage of a broad field of view to collect the ES cells, and changing the objective is not required. The general range for the number of ES cells that can be injected inside a blastocyst is considered to be between 10 and 20. The authors’ preference is to inject 15 ES cells, which balances the chance of germline transmission versus normal homeostasis in embryonic development. It is advisable not to inject more than 20 ES cells in that it is thought to be detrimental to the developing mouse embryo.
Note: The cells do not have to be touching each other when inside the injection pipette. If the cells are touching, there is a possibility that the cells may stick to each other, and many of the cells may come out of the blastocyst when withdrawing the injection pipette.
Note: The blastocyst can be oriented with the injection pipette by moving the joystick in either a clockwise or counterclockwise motion. Using the holding pipette to orient the blastocyst will disperse the cells away from the injection area and also other blastocysts.
Note: It is better to inject at the junction between the trophoblast cells, reducing damage to the embryo. Sometimes this junction may be hard to see.
Note: Touching the ICM is considered harmful to the blastocyst. If the injection pipette is inserted too slowly, the blastocyst will likely collapse, making injection difficult. If the pipette is only partially inside the blastocoel cavity and the blastocyst collapses, media can be injected to reexpand the blastocyst, and another attempt to inject the blastocyst can be tried. Otherwise, move the blastocyst to the side and perform injections on another.
Note: When injecting, consider the pressure inside the blastocoel cavity. If too much media is injected inside the blastocyst or too much holding force is applied by the holding pipette, the injected ES cells may be forced out of the blastocoel cavity.
Note: All blastocysts should be injected within a half hour of putting them on the injection dish, and the ES cell clones should be injected within 2 hours after putting them on the injection dish. If both pipettes are flushed out with oil, they have the potential to be reused for another injection day. Some people use a DNAse solution on top of the oil overlay to minimize stickiness due to cells and other debris.
Surgical transfers have to be performed the same day as that of ES cell injections, and E2.5-day-old pseudopregnant recipient females are generally used to allow for the blastocyst to catch up developmentally prior to implantation at E4.5–5.0 days. If E2.5 recipients are unavailable, the authors have had success with E1.5 and E2.0 recipients with this procedure. Pups are generally born 18 to 21 days p.c. of the pseudopregnant recipient. It should be noted that blue beads could be used as an alternative to bubbles when loading the transfer pipettes with embryos. The authors recommend a zoom setting of 3.2 for the surgical microscope, which will allow a large enough field of view of the uterine horn and still be able to the see the hole in the uterine horn wall.
Note: If an oil droplet forms on the outside of the transfer pipette upon placing it into the drop of media, gently shake the transfer pipette back and forth until the drop of oil comes off.
Note: The ovary and fat pad may be seen through the dorsal muscle if there is a low amount of fat on the mouse. The ovary will have a bright red color due to the formation of the corpus luteum. Do not use mice that lack a corpus luteum on the ovary. This would indicate that the mouse is not pseudopregnant. The ovarian fat pad will appear bright white in color with a smooth texture. Avoid the rough-looking, beige tissue that is attached to the spleen, which is most likely the pancreas.
Note: If the needle is inside the lumen, it should move freely with no resistance.
Note: If the bubbles inside the transfer pipette do not move when blowing them into the lumen of the uterus, the transfer pipette maybe touching the inside uterine wall. Slightly pull back the transfer pipette and try again. If this does not work, take out the transfer pipette and place in a dish containing a small amount of injection media under the dissecting microscope. If a blood clot is seen, dislodge it and reload the transfer pipette again with the injected blastocysts. Repeat step 18.
Note: The hole for the transfer pipette should be made near the tip of the uterine horn just as the taper from utero-tubal junction ends and becomes full size. This will allow for a higher rate of implantation than transferring near the cervix. There is also the added advantage of being easier than transferring at the very tip of the uterine horn especially since there is a much bigger lumen to put the 25-gauge needle and transfer pipette into.
Note: Enough embryos should be transferred to allow the embryos to be born healthy. If too few embyos are born, the pups will tend to be large in size, causing trauma to the pups and mother, as well as decreased lactation. If too many pups are transferred, the pups could be severely runted and die. Based on the authors’ experience using B6D2F1 and B6CBAF1 recipient mice, only 6 mice will be implanted per uterine horn; none of the blastocysts will migrate to the other uterine horn. At least 5 embryos should be transferred into each uterine horn with a maximum of 12 embryos transferred if transferring into both horns. If new to this surgical procedure, allow for a greater number of embryos when transferring to allow for enough embryos to implant.
C57BL/6 females at 3 weeks of age are considered to be the best choice for superovulation in the production of blastocysts. If not available, then 4-week-old females would be the next best choice. Mice 5 to 6 weeks of age or older are not considered the best choice for superovulation because of the fact that the females would be producing these comparable hormones themselves. If using C57BL/6 females at 5 to 6 weeks of age, you should not use hormones for superovulation and let the mice naturally ovulate instead. An advantage of superovulation is that fewer mice are needed to produce the number of blastocysts required for ES cell injections. The drawback with superovulation is the tendency to produce abnormal embryos. However, once at the blastocyst stage, most abnormal or defective embryos should have died or have stopped at an earlier developmental time point. C57BL/6 blastocysts are generally considered the standard in blastocyst injection to generate chimeric mice. However, other groups have used different strains for this purpose. The strain of mice chosen would depend on the ES cell line being used.
One hormone that we use in our lab is pregnant mare serum gonadotropin (PMSG), which is a follicle-stimulating hormone (FSH) involved with the formation of the egg follicles. Another hormone we use is human chorionic gonadotropin (HCG), which is a luteinizing homone (LH) that releases the formed eggs and also acts as an attractant to the stud male. Ten C57BL/6 females are used for each injection day, and 1 female is mated to each stud male. C57BL/6 males are only used once per week to ensure plugging and fertility. A light/dark cycle of 14 hours of light 10 hours of dark is a good choice for the rooms where the animals will be held. It is thought to be more conducive for mating.
Note: Actual times may vary depending on your animal facility and other variables. The plugging by C57BL/6 males becomes variable after 10 months of age with a reduced number of blastocysts and a greater number of earlier stage embryos. At about 11 months to 12 months of age, mice that are superovulated may produce only zygote-stage embryos at E3.5 p.c. At one year of age the C57BL/6 males will be become sterile. The C57BL/6 stud males are replaced at 9 to 10 months of age.
Various strains of mice can be used for the purpose of pseudopregnant foster mothers. Outbred mice tend to be the best mothers, and 2 notable examples are the Swiss Webster and CD1 strains. C57BL/6 hybrid strains are another alternative, as well as B6D2 and B6CBA. The use of inbred strains is generally not recommended, but if it is unavoidable, the FVB/N strain is the best choice. The strain of mouse chosen should be proven to have good mothers that take care of their offspring.
Vasectomized males of just about any strain could be used as long as they are known for their ability to plug females. Balb/c males seem to be the most commonly used for this purpose. Using strains of a different coat color can be useful to detect leakiness of the vasectomized male, allowing one to distinguish truly chimeric offspring from those of the leaky males.
Even if the vasectomized mice are from a reliable supplier, the males should be tested for sterility. Before the vasectomized males become too old and lose their plugging ability, new ones should be ordered and tested prior to use. The males could be mated for a 2-month period with females, or if time is a factor, there is the option of superovulating the females with PMSG and HCG.
Note: The authors initially mate 25 females 4 days before the start of ES cell injections. An additional 25 females are used to replace any plugged females. Unplugged females are kept with the males until 2 days prior to the last injection day for the week. Any unused females will then be separated out from the males and then can be reused for future matings. A period of at least 5 days should elapse before the mice are mated again. This will allow the mice to reestablish the estrus cycle.
Note: The time window from when the female is plugged by the male to when the plug can no longer be seen is thought to be 12 hours. If needed, females can be mated with the males in the morning to generate plugged pseudopregnant recipients in the afternoon.
Some of the pipettes used in the making of chimeric mice require a lot more skill and practice than others. The pipette that is the most difficult to make is the injection pipette followed by the transfer pipette and then the holding pipette. Three variables that will affect the quality of pipettes produced are the melting of the glass tubing, the length of time the glass tubing is pulled, and how much force is applied to the glass tubing during the pull. When first learning to pull the transfer pipette under a microflame, these variables are harder to control. Experience is required to control these variables in being able to pull a good transfer pipette. For the pipette puller that the authors use in the lab, time and force of the glass tubing is controlled by the solenoid, and temperature is set by the heater control knob. How fast the ends of the pipettes are melted by the glass bead on the microforge will depend upon how close the pipette is to the glass bead, the heat level setting of the filament, and the size of the glass bead.
Both injection and holding pipettes can be purchased from vendors, but the expense of these premade pipettes can be costly if a lot of them will be used. After the initial expenditure of buying the equipment, such as a microforge and pipette puller, the real expense is that of the glass tubing.
Note: The pipettes can be broken right after the pipettes have been pulled and allowed to cool for a few seconds. A diamond pencil can be used to score the glass first before breaking it. Another alternative is to break the pipettes between the thumbnail and forefinger.
Note: Viewing the pulled transfer pipette takes practice. The only way to gauge the size of the transfer needle is to use a microforge that has a reticle in the eyepiece.
Note: Avoid any areas that are chipped, broken, or cracked.
Note: The inner diameter size of the transfer pipette should be big enough so the fully expanded blastocysts are not constricted, which could possibly damage them. A pipette too small is more susceptible to blockage by blood clotting. The pipette should also not be too big that the blastocysts are bunched up together side by side instead of being lined up in a row. A general range for the inner diameter would be approximately 8 to 10 units. If using a reticle to make different types of pipettes for ES cell injection, the measurement scale for the reticule at 40× magnification is 1 unit = 2.5 μm, and at 100×, 1 unit = 10 μm.
Note: For the inner diameter of the holding pipette, it is necessary to use a higher magnification than 40× to make the holding pipette inner diameter of 20 μm. Magnification of at least 100× will work well for this purpose. If 40× magnification is used, the size of the inner diameter will be inconsistent and most likely will be larger than 20 μm. Inconsistency in size will lead to control problems with the holding pipette. This is especially important if the blastocyst is held by the trophoblasts and not the inner cell mass during injections. This can make injections very difficult, and the blastocyst could be suctioned into the holding pipette. The authors use a heat setting of 12.0 units and a solenoid setting of 1.0 A for pulling holding pipettes.
Note: When the bevel is facing toward you to the side, it is easier to see the edge of the 30-ga needle.
Note: With the settings that authors use for pulling injection pipettes, the pipette can be broken a few times to get a proper broken pipette. After several attempts, it is best to use another pulled pipette.
Note: For the inner diameter of the injection pipette, we prefer a size of ~20 μm.
Note: The temperature setting should not be too high or the inner diameter will become smaller. The temperature of the glass bead should be high enough to actually melt the pipette tip. Melting and pulling the pipette tip too much will create a bevel that is too long, which would make ES cell injections more difficult. If the tip is too long or there is a filament like structure on the tip, the tip can be gently broken again and the melting of the tip can be repeated. If the tip is still is too short after the first melting it can be repeated. If the inner diameter becomes too narrow or has a bevel too long, it is best to start over with another broken injection pipette. The authors prefer an injection pipette with an inner diameter of approximately 20 μm, which can be successfully obtained by using a heat setting of 14.2 units and a solenoid setting of 4.2 A when pulling injection pipettes.
Various strains can, in theory, be used as the host blastocyst and as the ES cell line. Generally the most common strain of mouse used for the host blastocyst is C57BL/6 mice and 129 strain ES cells. It is best to use strains of mice that are different in coat color for the blastocyst and the ES cell line. The mating scheme will be determined by what strains of mice are used in the generation of chimeric mice.
If one is using a C57BL/6 donor strain for the blastocyst, the chimeras could be mated with either a C57BL/6 or a Black Swiss to generate germline mice. Each of these mice has disadvantages and advantages. The Black Swiss mice are better breeders, and the mothers take better care of their offspring and, unlike C57BL/6 females, are not disturbed easily by noise and vibration. However, if C57BL/6 mice are used, the genetic background is conserved, and if Black Swiss mice are used, this might be lost since they are outbred mice. This could have possible implications on the mouse phenotype for the knockout mouse created. Also, if using either the C57BL/6 or Black Swiss strains of mice to mate with the chimeric mice, the germline mice will be on a mixed strain background, and in order to achieve an isocongenic C57BL/6 or 129 strain background, the mice will have to be backcrossed at least 10 times. It is possible to use speed congenics and PCR methodology to limit the amount of backcrosses needed for a deleted gene of interest on a particular isogenic strain background.
Another option is the mating of the C57BL/6 x 129 chimera with an isogenic strain of 129 mice that has been used to generate an ES cell line. The 129 isogenic strain can be mated to the chimera to obtain F1 founder mice on the isogenic strain of the ES cell line in only one backcross generation. One drawback to this scheme is that some strains of 129 mice have low fecundity and may result in breeding problems in the analysis of the mouse knockout phenotype. If ES cells have contamination from other strains in the genetic background, then the mice will still have to be backcrossed at least 10 times in order to obtain isocongenic purity.
If using a color marker to determine the chimeric composition of the ES cells, it is best to choose the chimeras with the highest contribution possible. They should be robust and preferably male, and they should not be dwarfs. Female chimeras could be used if needed, but are not preferred in that ES cells are generally male. Any female chimera would likely have lost the Y chromosome, becoming XO, and this is thought to be more unstable.
Female chimeras have been known to generate germline mice even with the loss of the Y chromosome. Another advantage of using male chimeras over females is that a large number of offspring can be generated in a short period of time, ensuring the transfer of the knockout allele to the F1 generation offspring.
When mating the chimeras, the most efficient mating scheme would be the mating of 2 females to 1 male chimera, Another approach is the so-called “shotgun” approach, the mating of 4 females to 1 male chimera. One drawback to this approach is that there could be a delay in the generation of offspring because of the fact that the males may become distracted by so many females and delay plugging. Furthermore, there could be a lot of mice generated at one time, which would take up a lot of cage space.
The mating scheme chosen to generate germline mice must conform to the ACUC regulations of the institution where the animal work is taking place.
Use tissue culture-grade reagents when working with cells and embryos. For common stock solutions see APPENDIX; for suppliers see SUPPLIERS APPENDIX.
To make 100% Avertin stock solution, add 50 ml of tertiary amyl alcohol (Fisher Scientific A730-1) to 50 g of 2,2,2-tribromoethanol (Fluka catalogue # 90710). Add the smallest stirring bar possible to the mixture and place on a stirring plate. Not all 50 ml of tertiary amyl alcohol may fit inside the 2,2,2-tribromoethanol bottle due to the stirring bar inside. Allow the mixture to stir at room temperature until the stirring bar is heard. Keep stirring the mixture overnight at 4 °C to make sure the 2,2,2-tribromoethanol is dissolved into solution. The next day with a 1 ml pipette, check to see if the 2,2,2-tribromoethanol is completely dissolved. If there are no solid crystals in solution, the stock solution is ready. The color of the Avertin stock solution should have a clear to very pale yellow color. If the color of the solution has a dark brown color, discard solution. For the 2.5% working solution, allow sterile 1× DPBS to warm up to room temperature. To 10 ml of DPBS add 250 μl of Avertin stock solution, and shake and vortex until fully dissolved. Filter this solution through a 0.22-μm, nonpolystyrene filter unit. Store in a polypropylene tube covered with foil.
There will be some variation in each batch of stock Avertin, so it is essential that it be tested on mice prior to using in surgery for efficacy and toxicity. Both solutions should be kept to a minimum at room temperature and shielded from light, which will cause the breakdown of Avertin into toxic products. If a glass container is unavailable, use plastic containers such as polypropylene. Avoid using polystyrene, as it will chemically react with the Avertin stock solution and become toxic to the mouse and may lead to peritonitis. The recommended dosage of Avertin is between 0.015 and 0.017 ml/g body weight. If the working solution is being reused, it is suggested that the solution be refiltered soon after use to reduce the risk of possible bacterial contamination. If it is thought that the working solution is contaminated, discard it.
Under sterile conditions, add 10 ml DPBS to a vial containing 10,000 units of chorionic gonadotropin (Sigma #CG10). Take 2 ml of this solution and dilute into 40 ml of PBS. Aliquot this solution into tubes and store in a −20 °C freezer.
81 ml Dulbecco’s Modified Eagle’s Medium (DMEM) containing 4,500 mg/L D-glucose, L-glutamate, and 25 mM Hepes (Invitrogen #12430054)
8 ml of fetal bovine serum (preferably already ES cell tested)
0.5 ml penicillin/streptomycin containing 10,000 units per ml penicillin and 10,000 μg/ml streptomycin (Gibco #15140-122)
Filter the prepared media with a 0.22-μm filter unit. Divide the injection media into three 50-ml tubes. Use one tube for each injection day.
Place 30 μl of injection media on a 3-ml tissue culture plate. Overlay the drop of media with prefiltered, embryo-tested quality mineral oil (Sigma #M-3516 or equivalent).
An inverted microscope is generally used, and it should have 10× and 20× objectives, with the 40× objective being optional. The 20× objective is more useful for those who are first learning ES cell injections, but the 10× objective is used more often by more experienced researchers. There are 3 optic systems most commonly used for microinjections. Hoffman generally works best for viewing items through plastic, Nomarski or differential interference contrast (DIC) is better suited for looking through glass, and phase contrast can be used for either plastic or glass. In the authors’ lab, DIC optics are used in the microinjection microscope. General injection and holding line systems would consist of a syringe-type unit, line tubing, and pipette holder and may or may not have a stopcock attached to an oil-filled syringe. Luer locking or other types of connection systems should be used whenever possible to minimize or eliminate possible leaks that might occur. For the injection and holding line systems, one that is filled with oil, rather than air, will tend to give better control. Silicone oil would be a better choice than mineral oil in that mineral oil will tend to shrink, causing air bubbles that affect control. An injection line system with a stopcock and a tuberculin-type syringe with an oil-filled reservoir will allow the flushing out of debris and air bubbles from the injection and holding lines and both pipettes. A 12-cc oil-filled syringe reservoir will allow the microsyringe units to be aligned back to the midpoint, as opposed to one that does not.
For a microsyringe controlling unit, the authors prefer a device made by Minnetonka Instruments, which has an internal Hamilton syringe and an external Mitutoyo syringe. The injection line tubing and Hamilton syringe should be a smaller volume than the holding system because the injection pipette is much smaller than the holding one. As for micromanipulators, the authors have had successful experience with Narishige and Leica manual manipulators. There are multiple options available, and users should decide which model will work best for their use. It should include fine as well as coarse movement. A manual model is preferred for the actual injection of the blastocyst, because it allows for more precise movement and control, unlike some powered systems. To avoid any vibration problems that could affect microinjections, an air suspension table using N2 or CO2 gas cylinders should be considered.
Under sterile conditions, add 3 ml of sterile PBS to a vial of 2000 units of PMSG (Sigma #G 4527). Add this solution to a tube containing 40 ml of PBS, aliquot it into tubes, and store the tubes in a −20 °C freezer.
Add 1.25 ml of Tween 80 solution (Fluka #93780) into 98.75 ml of sterile H2O. Filter through a 0.22-μm filter unit.
Early cancer studies in mice initially done by (Stevens 1970, 1973) led to the generation of the strain of mice known as the 129. Subsequent developmental studies in these mice made possible the creation of 2 main mouse cell lines, the embryonal carcinoma (EC) cells and the embryonal stem (ES) cells. EC cells were originally derived from teratomas that came from Stevens’s mice (Kahan et al. 1970; Rosenthal et al. 1970). ES cells were initially derived from the inner cell mass (ICM) of mouse blastocyst embryos from 129 mice (Evans and Kaufman 1981; Martin 1981).
With the development of the ES cell injection technique into blastocyst mouse embryos (Gardner 1968), the true potential of these cell lines to be able to contribute to the developing mouse embryo could then be assessed. The EC cells initially showed promise with pluripotency in the formation of chimeric mice and the contribution to various tissues (Brinster 1974). However, EC cells had problems with developmental abnormalities, tumors, and a lower germline transmission efficiency compared to ES cells (Papaioannou and Rossant 1983). ES cells, on the other hand, showed this pluripotency in chimeric formation and totipotency in germline transmission to the offspring without the problems associated with EC cells (Bradley et al. 1984).
Upon understanding the mechanisms of homologous recombination, the possibility of generating knockout mice became possible. Folger et al. (1982) were the first to demonstrate that through homologous recombination, nonreplicating DNA could be transferred into a mammalian cell. This would later lead to “gene targeting” of a plasmid into the β-globin gene of mammalian cells (Smithies et al. 1985). The first gene to be targeted in ES cells was the hypoxanthine phosphoribosyltransferase (Hprt) gene (Thomas and Capecchi 1987; Doetschman et al. 1988). This had the benefit of it being X-linked, and only one copy was needed to show deletion in male ES cells. Later, ES cells with the Hprt mutation would possess the ability for germline transmission (Hooper et al. 1987; Kuehn et al. 1987; Koller et al. 1989; Thompson et al. 1989). While the generation of a mutant mouse with this gene showed initial promise as a model for Lesch-Nyhan syndrome, the phenotype did not correlate with the human disease (Samuel et al. 1993). The Hprt knockout mouse phenotype highlights what may happen when one attempts to delete a human gene in the mouse.
An alternative approach to generate chimeric mice is the ES embryo aggregation technique. This technique can be further divided into 2 basic methods. One of them is the diploid aggregation technique, which involves ES cells cultured with morula stage embryos. This can further be divided into 2 techniques. One technique developed by (Wood et al. 1993) involves culturing the morula on a layer of ES cells. The other method, devised by (Khillan and Bao 1997), also uses morula and ES cells but with a defined microwell and a given number of cells. A second morula aggregation method, in contrast, uses a tetraploid developed by (Nagy et al. 1993): 2-cell embryos are electrically fused together and cultured until the 4-cell stage. These 4-cell tetraploid embryos are then used to “sandwich” the ES cells, allowing integration (fusion) to occur. This is done in a well on a tissue culture dish. All 3 methods have the advantage of not requiring injection skills or the expensive equipment needed to perform the ES injection procedure.
The last key component to generating chimeric mice and subsequent knockout mice is the uterine surgical transfer technique. This technique was initially developed by (McLaren and Michie 1956). They determined that a critical parameter for success in the uterine transfer technique is the surgical transfer of E3.5 blastocyst embryos into E2.5-aged recipient females. Based on this work, (McLaren and Biggers 1958) cultured morula-stage embryos in vitro to the blastocyst stage and surgically transferred them by the uterine technique to successfully generate live offspring.
A recent development suggests the possibility of using other types of stem cells instead of ES cells to generate chimeric and knockout mice. Guan et al. (2006) isolated spermatogonia stem cells (SSC) and maintained them under ES cell growth conditions. As a result, these cells retain characteristics of both SSC and ES cells, and they were subsequently named multipotent adult germline stem cells (maGSCs). This technique may be another method to derive cells that are ES cell capable, without having to obtain them from the inner cell mass of a blastocyst. This may make it easier to acquire ES-like cells from a strain of mouse where currently none is available.
There are 3 main areas where problems may be encountered while creating chimeric mice: harvesting blastocyst embryos, injecting ES cells into the blastocyst embryo, and surgically reimplanting embryos into pseudopregnant female mice. The ES injection technique can largely be controlled by conditions in the laboratory, which can make problems in this area easier to solve. The other 2 areas of harvesting the blastocyst embryos and surgically reimplanting them depend on conditions in the lab, the in vivo variability of the mice themselves, and conditions in the animal mouse facility. Thus, solving problems in these last 2 areas can be quite complex. It is probably best to look at laboratory conditions first, which are more easily controlled, then proceed to looking at the mice themselves and to the animal facility conditions. It is important that detailed notes be taken, which can greatly help to pinpoint problems. Aspects of all related procedures should be assessed periodically so that potential problems can be addressed early.
The number of blastocysts obtained through the process of superovulation and harvesting of blastocyst embryos can depend on several factors. The age of the mice and the conditions in the animal facility are 2 areas affecting blastocyst yield. Females that are 3 weeks old are preferable because they are more easily induced by the PMSG and HCG hormones. If not available, 4-week-old females are the next best choice. Female mice 6 weeks or older should be producing their own reproductive hormones, which could interfere with any hormone injected intraperitoneally. For the breeding stud males, a range of 7 weeks to ~ 10 months of age has worked well in our experience. Around 10 months of age, plugging by the stud males becomes variable. Another sign that the stud males are too old is that only very early stage embryos are recovered. Replacing the stud males will solve both of these problems. In the animal room where the mice are being held, the most important factor is the timing of the light and dark cycle. A light and dark cycle of 14 hours light and 10 hours dark is considered to be conducive for the mice to mate. The light cycle should be checked if there is any doubt that it is working correctly.
Hormone-related issues can also influence blastocyst yield. These would include the source where the hormones came from, a possible bad lot, or how the hormones were prepared. A reputable source that others have had success with should be the first choice. Another lot number of the same hormones should be tried to see if similar results are obtained. If the hormones have not been diluted to the proper concentration they should be discarded. Freezing and thawing of hormones is to be avoided as this will decrease the activity and effectiveness. The proper dosage of hormone being injected intraperitoneally and the timing of the hormone injections are also important.
What day and time the blastocysts are to be harvested will determine the timing of the hormone injections. If at the desired time of day only morula or earlier stage embryos are obtained, both of the hormone injections should be done at earlier times. If only hatched blastocysts are collected, hormone injections should be done at later times.
When new to the technique of harvesting and collecting blastocysts, it is not uncommon to obtain a lower embryo yield than that of someone who is experienced. Blastocysts can be lost while harvesting the uterine horns. Since the blastocysts are free-floating inside the uterine horns, care should be taken to not cut holes in the uterine wall when cutting off the mesometrium structure. Letting the uterine horns snap like a rubber band should also be avoided when extracting out the uterine horn as this can also contribute to losing blastocysts. After all of the uterine horns have been taken out of the original holding dish, check to see if there are any blastocysts left on the dish. If there are a lot of them on the dish, then there is likely a problem with the harvesting technique. Practice is needed until the technique is mastered. The way the uterine horns are flushed out can also affect blastocyst yield. The opening of the cervix should be oriented slightly up and to the side. This will allow one to see the injection media being flushed out of the cervix while keeping the flushed media inside the 3-cm dish. In our experience, a 30-ga needle works best for flushing the blastocysts out of the uterine horns. A good amount of pressure can be applied to the syringe with less chance of media coming out of the dish, unlike that of larger gauge needles. It should be noted that even under optimal conditions the process of superovulation and harvesting the blastocysts doesn’t always work well all the time. There will be days when very few embryos are recovered.
Blastocysts are very hardy embryos and can withstand injections quite well, unlike embryos at earlier developmental stages, such as zygotes. There are 3 main areas where problems might arise during ES cell injections of blastocysts: the injection and holding line systems, the injection pipette, and the holding pipette.
In the injection and holding line systems, problems that may commonly occur are that the oil does not move inside the pipette, there is a lack of control or the movement is very slow, or the cells are continuously taken up by the holding and injection pipette when the line is open. These problems could be due to air in the system, there maybe a leaky or worn-out stopcock, or there could be a leaky joint or connection. If air is found in the system, a 19-ga needle and syringe filled with oil (or the oil-filled syringe reservoir) is a good way to purge this out. A leaky stopcock due to wear must be replaced with a new one. Reconnect the loose joint or connection and replace any fittings or pieces of tubing if necessary.
Another area where problems might arise could be the injection pipette. Debris or oil can become stuck to the inside and outside of the injection pipette. There maybe blockage or air bubbles that form inside the injection pipette. Cells may lyse inside the injection pipette. The injection pipette may not penetrate inside the blastocyst and collapse before penetrating the blastocoel cavity. Debris, silicone oil, air bubbles, or other types of blockage may be flushed out with the syringe reservoir filled with silicone oil. If this method does not remove the debris or oil attached to the inside of the pipette, a smaller MEF cell maybe used much like a pipe cleaning device. Debris on the outside of the injection pipette may be removed by moving up the injection pipette through the media/oil and oil/air interfaces. The holding pipette could be used to pull debris off the injection needle, but care should be taken not to block the inside of the holding pipette. The reason for lysis of ES cells inside the injection pipette could be due to the fact some of the ES cells inside the pipette are too large, of poor quality, or have come in contact with oil. ES cells should be well typsinized to remove cell surface proteins and kept cold to impede them from reforming. If not, this can cause “stickiness” inside and outside the injection pipette, impeding injections. If the inner diameter of the injection pipette is too small, then the cells could lyse. Failure to inject into the blastocoel cavity can be the result of a dull or badly damaged injection pipette. It is best to inject between the junction of the trophoblast cells, which will make injections easier. The injection pipette should be at approximately the midpoint of the blastocyst height so that it will be in the center of the blastocoel cavity. If it is on the bottom of the blastocoel cavity, it will be harder to inject. The injection pipette should be raised up to avoid this from happening. It is very important to have really good injection needles when new to the injection procedure as this will make injections much easier and can help avoid much of the frustration that is often encountered when learning the injection technique. If all other attempts fail to correct problems encountered in injections of the blastocyst, the injection pipette needs to be replaced.
The third and last area where problems can occur is that of the holding pipette. It could become blocked with debris or have air bubbles forming inside. Erratic holding control may be experienced or there is no holding suction at all. If bubbles form or the inside of the holding pipette becomes blocked, the holding pipette may be flushed out with the syringe reservoir filled with silicone oil. Usually this method will not work with a large blockage. If this happens, the holding pipette will need to be replaced. In our experience, a holding pipette with an inner diameter of approximately 20 μm works best for the holding control of the holding pipette. A smaller diameter may not have enough holding power, while a larger diameter tends to give more erratic control and the possibility of the blastocyst ending up inside the holding pipette. If the blastocyst moves when attempting to inject, this could be due to the fact that the holding pipette is too low to the surface of the well of the injection dish and there is not enough room between the holding pipette and the surface of the well. It also may mean that the holding pipette is not properly aligned or is too close to the shoulder of the injection dish. Touching the blastocyst to the bottom of the well adds much-needed stability for holding.
There could be a number of reasons for a small litter size or for why pseudopregnant females are not taking care of the pups. The best place to start is to do in utero analysis of uninjected blastocysts by extracting out the uterine horns of pseudopregnant recipient females and looking at E9.5 embryos. If there is a low percentage of embryos being implanted, this could most likely be due to factors related to the surgical technique.
Surgical technique can be influenced by several factors. Only perform the uterine surgical transfer procedure on mice that have the corpus luteum (red spots) present on the ovary. Absence of the corpus luteum suggests the female was not plugged or was not in estrus when plugged by the vasectomized male. Avoid touching the uterine horns and ovary when possible, except to hold the uterus to make the hole inside the uterus to the lumen and inserting the transfer pipette. When making the surgical incision in the dorsal muscle wall, avoid cutting the blood vessels, which can cause excessive bleeding making it hard to find the ovary and fat pad. Like the surgical incision, avoid the blood vessels in the uterine wall when inserting the 25-ga needle in the uterine wall to make the hole in the lumen. If a blood vessel is damaged during this procedure, bleeding might clog the inside of the transfer pipette. Mouth pipetting the embryos will be difficult if not impossible. Reload the transfer pipette, and make another attempt to transfer the blastocysts. Both the 25-ga needle and transfer pipette should pass freely inside the hole in the uterine wall. This indicates that both have entered inside the lumen. The attempt to transfer the blastocysts will not be successful if there is any obstruction. The procedure of mouth pipetting, the quality of the transfer pipette, and how the transfer pipette is loaded can influence the outcome of the surgical transfer procedure. As the investigator becomes skilled in using the mouth pipette, this factor can largely be ruled out. If the inner diameter of the transfer pipette is too small, there is a tendency for the transfer pipette to become easily clogged by bleeding that can happen during the procedure, and if the inner diameter is too big, this may affect the control in that it will be too sensitive and the blastocyst will migrate up the transfer pipette. Also, a transfer pipette that is too small in diameter may make it difficult to blow the blastocysts inside the lumen of the uterus. The way a transfer pipette is loaded with blastocysts can make a difference in the transfer results. There is a lot of variation in the way a transfer pipette can be loaded with blastocysts. Air bubbles and oil are factors that can help to stabilize the embryos inside the transfer pipette, but will move when blown out with the mouth pipette. After the transfer of the blastocysts has taken place, examine the transfer pipette under a dissecting microscope to see if there are any left inside. If any blastocysts are left, they should be transferred again until they are no longer in the transfer pipette.
If the rate of implantation of E9.5 embryos is high, then there may be other factors other than the surgical technique, one of these being related to the culture conditions of the ES cells and blastocyst embryos. Reagents that are used to grow the ES cells or that may come into contact with the blastocysts could be the culprit. Serum, DMEM media, and oil should be tested on the parental cell line first before they are used on targeted ES cell clones. Reagents used more specifically with the in vitro conditions of the blastocyst should be tested before they are to be injected with ES cells. If these reagents have been tested before, they are most likely not the cause. However, if a new lot is now being used, it is worthwhile to investigate. Another lot should be tried to see if similar results are obtained. If a new lot of a particular reagent has been determined as the cause, it should be discarded and replaced as soon as possible. Haploid insufficiency of the targeted gene or the way the construct is designed may influence embryo lethality and pup death; if other targeted ES cell clones are generally doing well, then this could be the cause.
The other area that may affect the gestation of the pseudopregnant recipient litters is the conditions in the animal room where the mice are being housed. Care should be exercised not to stress the recipient mother because it could cause physiological conditions leading to reabsorption of the mouse embryos in utero and the death of the newborn pups. Probably the key embryonic developmental time points where this stress would likely be the most detrimental are at E4.5 to E5, when the embryos implant, and shortly before birth. The first week of birth is the most critical time for the newborn pups. If they survive this week, there generally should be no complications. Stress factors to avoid include the Bruce effect, ammonia levels, noise, vibration, animal handling, animal food nutrition, ventilation, and the light/dark cycle. These conditions in the room should be monitored to determine which of them might be the cause. Infectious agents such as viruses, which may be present as a latent infection in the animal colonies, can affect knockout mouse production.
Finding the exact cause of embryo lethality can be difficult and time-consuming as numerous factors affecting pregnancy need to be ruled out. Even the simplest of factors might be the cause of the problem and should not be overlooked. For example, we determined that a defective wheel on one of the animal racks was the possible cause for the pups being reabsorbed in utero and the pseudopregnant mothers not taking care of the pups after birth. When the rack was replaced, these problems went away. Lastly, one factor that is often overlooked is the effect of lighting on embryos, whether it is in the lab or surgical room. This could cause apoptosis to occur in the ICM of the blastocyst embryo, which would have detrimental effects on injected embryos and the potential for live pups being born.
With practice, this surgical transfer technique should result in at least 70 to 80 percent or better of the surgical recipients giving birth. Even uninjected blastocysts, when surgically transferred and implanted, may end up being reabsorbed during gestation. This can happen as early as E9.5. The number of pups born will be influenced by the strain of female mouse used for the pseudopregnant recipient. In our experience, the B6D2 and B6CBA strains of mice have only given birth to no more than 6 pups per uterine horn, regardless of how many were surgically transferred. If more than 6 embryos are surgically transferred, these extra embryos most likely will not implant.
Blastocysts are very hardy embryos, especially when compared to early stage ones. They tolerate changes in temperature, pH, and the technique of ES injection. Unless there is severe trauma of the embryos, like that of damaging the ICM, the embryos should not be negatively impacted. In the hands of a skilled injectionist, the ES cell injection procedure should not adversely affect the birth rate of pups being born.
There are numerous ES cell lines available for the purpose of making chimeras to generate knockout mice. Testing of the parental ES cell lines the investigator plans to use should be first evaluated for any possible ill effects from these cells. Their efficiency in generating chimeric mice needs to be evaluated before carrying out any gene targeting related experiments on them. Some markers to look at in evaluating the effectiveness of the parental cell include the percentage of chimerism, the ratio of the number of male chimeras born to females, and the ability to generate germline offspring. In a good parental ES cell line, the chimerism should be high, the male-to-female ratio high, and the number of pups needed to see germline transmission low. It is possible to see chimerism or germline transmission as early as 5 days with a small patch of fur becoming visible. This is much easier to see if an “albino” ES cell line is used, in that these white areas would appear nude before the fur starts to grow in. Germline transmission on other types of 129 ES cell lines used with a C57BL/6 host blastocyst will have either a medium brown or agouti (golden brown) appearance.
Even if good parental lines are used, there can be great variability in a targeted clone’s ability to create a high percentage of chimeras and in the germline potential of these chimeric mice. This is most likely due to trauma the ES cells have to undergo in the electroporation of the gene targeting construct and selection process to determine a correctly targeted clone. In theory it should take only 2 clones to generate 2 different germline chimeric lines leading to the establishment of 2 independent knockout mouse lines. This would confirm that the phenotype is due to the gene targeting construct and not to the cells themselves. Several targeted clones should be available to ensure that there is germline transmission from 2 independent clones, should some of the clones give low chimerism and are not germline.
A situation may arise where lethality is most likely related to the clones being injected or the construct that is electroporated inside the ES cells. This can result in no chimeric pups being born at all or only a few surviving to birth and then dying shortly thereafter. If everything is generally going well with other constructs and ES cell clones, then other factors affecting gestation can be ruled out. If the ES clones in question have been injected additional times and large numbers of blastocysts have been injected with similar results, then this further suggests a construct-related problem. Further injections will most likely be of little use. There is also the possibility of embryonic and perinatal lethality occurring in the germline F1 mice. As previously mentioned with chimeras, this could be due to targeting construct-related issues. The construct should be reevaluated and any problems corrected. Additional clones can be generated and injections performed. Lastly, lethality could be due to the function of the gene deleted and its essential role in embryonic development. This haploid insufficiency may be the reason for embryonic and pup lethality. If this occurs, the only kinds of phenotypic analyses that may be carried out would be in utero analysis of the developing embryos or in vitro tissue culture analysis.
Plan to start the initial intraperitoneal PMSG hormone injections 1 week before the day when the blastocysts will be needed. The entire process of harvesting the blastocysts, injecting them, and surgically transferring them back to a pseudopregnant recipient female is generally done the same day. Even for the experienced injectionist this can make for a long day, especially when problems with the injection system and bad cell morphology of the ES cell clones happen on the same day. The surgical procedure should be practiced before attempting the injection procedure since it generally takes longer to master. Having a video display for all the procedures aids greatly in the visualization of these techniques and can shorten the time required to learn them, which is expected to take 6 months or less. How long it takes will ultimately depend on previous experience, the person instructing the procedures, and the desire of the investigator to learn the procedures.