Recovery of Preimplantation Mouse Embryos from the Oviduct and Uterus

Summary

The primary role of the ovary is to generate viable eggs, which can be assessed by their ability to undergo fertilization and support preimplantation embryo development.

Abstract

One of the primary functions of the ovary is to produce healthy eggs for reproduction. One method to assess ovarian and defective reproductive tract function is by measuring fertilization capacity and preimplantation embryo development in the oviduct and uterus. In this study, female mice at estrus were mated overnight with a proven male, and the presence of a copulatory plug the next morning was recorded as 0.5 days post coitum (dpc). By 09:30 h on 0.5 dpc, both oviducts were dissected to isolate cumulus-oocyte complexes. Zygotes were identified at the pronuclear stage. The oviducts were then excised at 1.5 dpc and 2.5 dpc to retrieve 2-cell and 4- to 8-cell embryos using a blunted 30-G needle filled with Leibovitz-15 medium plus 1% fetal bovine serum. At 3.5 dpc, both the oviduct and uterus were flushed for morula and blastocyst stage embryos. The total number of fertilized eggs and embryos at each stage can be expressed as a percentage of the total ovulated eggs, serving as indicators of egg competency.

Introduction

The key roles of the ovary are to produce steroid hormones and healthy eggs for reproduction. Primordial follicles contain oocytes that are arrested in prophase I of meiosis I. Oocytes remain arrested in this state through follicular activation and growth. Oocytes are often considered to be in an incompetent stage due to their inability to undergo fertilization. In response to a luteinizing hormone (LH) surge, meiosis resumes, and the oocyte enters metaphase II (MII), now called an egg because it is competent for fertilization. The MII eggs (with the presence of the 1^st polar body) are mature but arrested at this stage until fertilization¹. Follicular development is tightly regulated by endocrine hormones through the hypothalamic-pituitary-gonadal (HPG) axis. When perturbations occur in the HPG axis, it alters ovarian physiology, which can result in dysfunctional follicular and oocyte development. In addition to measuring ovarian hormones (estradiol and progesterone), assessing egg fertilization capacity and preimplantation embryo development can be used to investigate ovarian function in vivo.

The oviduct is the location of gamete and embryo transport, fertilization, and preimplantation embryo development. As such, preimplantation embryo collection can be used to assess the reproductive consequences of pathophysiological, pharmacological, or genetic perturbations within the female reproductive tract. Our research projects often investigate preimplantation embryo development after genetic ablation of genes of interest in the oviductal epithelial cells. For example, we show that deletion of either the estrogen or progesterone receptors in oviduct epithelial cells results in embryo death prior to the 2-cell stage² or delayed preimplantation developmental outcomes³, respectively.

In mice, estradiol levels peak at proestrus, and females are receptive during estrus. Females are paired with the male overnight at the estrus stage. The following morning, the copulatory plug is visible and is considered 0.5 days post coitus (dpc). Eggs are ovulated as cumulus oocyte complexes (COCs), and sperm travel through the reproductive tract and fertilize the eggs in the ampulla region of the oviduct. Successful fertilization can be determined by the presence of 2 pronuclei and the 2^nd polar bodies at approximately 11:00 h of 0.5 dpc. At this stage, the embryos are called zygotes or 1-cell embryos. Preimplantation developmental trajectory can be further evaluated by the presence of 2-cell embryos at 1.5 dpc and 4- to 8-cell stage at 2.5 dpc in the oviduct. At 3.5 dpc, embryos develop to the morula and blastocyst stage and are now transported to the uterus. If embryos show delayed development or are retained in the oviduct at 3.5 dpc, it would result from poor-quality eggs (ovarian deficiency) or an altered oviductal environment. In this study, embryos were collected at each time point and evaluated using light microscopy.

Protocol

All animals and procedures used in this study were handled in accordance with University of Missouri (MU) Animal Care and Use Committee (ACUC) guidelines and MU-approved animal protocols. Control, progesterone receptor floxed (Pgr^f/f), mice and mice with a conditional deletion of Pgr in the epithelial cells of the reproductive tract (Pgr^d/d) are used in this study³. All procedures are performed without anesthesia. Use female mice needed for the desired experiment. To assess estrus and mating receptivity, a rapid (no staining) vaginal cytology evaluation is performed⁴. The reagents and the equipment used are listed in the Table of Materials.

1. Assessing estrus in female mice

1. At 11:00 h, hold the mouse at the base of the tail with the index finger and thumb to lift the hind legs, then use the third, fourth, and fifth fingers to press down on the back near the hip area (Figure 1A).
2. Dispense and aspirate ~40 µL of sterile saline into the vagina a few times. Do not insert the tip of the pipette into the vaginal cavity; only place the tip of the pipette at the vaginal opening while aspirating.
   NOTE: The pipette tip should remain at the vaginal opening throughout both dispensing and aspiration. The pipette tips should be sterilized prior to use (i.e., autoclaving), and a new pipette tip is used for each mouse. Do not reuse.
3. Put one droplet (~20 µL) of the collected vaginal wash on a slide. 6–8 drops will fit on a slide; therefore, samples from 6–8 mice can be collected and placed on the same slide (Figure 1A, bottom panel). But be careful not to jostle or mix the drops.
4. Using a 10x objective on a light microscope, evaluate the droplets for the types of cells present (Figure 1B).
   NOTE: It is not ideal for the droplets to dry, but they can be rehydrated by carefully adding saline before evaluation.
5. Pair only females at the estrus stage, as determined by the presence of enucleated cornified cells (Figure 1B), with the male mice.

2. Mating and accessing the copulatory plug

NOTE: Once the estrus stage is confirmed, follow the steps below to pair the mice and ensure that they mate. Use a wild-type male that is a proven breeder to pair with the females. C57BL/6J or B6D2F1/J males between 2–10 months are often used as the male breeder. If CD1 females are used, they should be paired with CD1 males due to the large body size compared to other common inbred strains.

1. At 16:00 h, house a single female in estrus with a single male overnight (Figure 2A).
2. At 08:00 h the following morning, evaluate the vaginal opening, holding the mouse by the base of the tail and lifting the hind legs, for an opaque-white semen coagulum (copulatory plug or plug, Figure 2B).
   NOTE: Often, the plug will be visible externally, and the external opening of the vagina (vulva) will appear large and swollen⁵. However, if the plug is not readily visible, an autoclaved blunted wooden toothpick can be used to gently open the vagina. If there is a plug, there should be resistance from a solid plug, and the end of the stick will not go deep into the vaginal canal (Figure 2B).

3. Preparation for tissue and embryo collection

NOTE: Now that mating has been confirmed, embryos can be collected at the appropriate time point. Usually, embryos are collected at 09:30 h of 0.5, 1.5, and 2.5 dpc for consistency. For morula and blastocyst collection, the embryos should be in the uterus at 13:00 h of 3.5 dpc.

1. Before the collection time, combine Leibovitz-15 media with 1% fetal bovine serum (L15 + 1%FBS). Sterilize the media by filtering using a 0.22-µm size membrane and place in a heat block set to 37 °C.
   ​NOTE: L15 media does not contain sodium bicarbonate, but is buffered using phosphates and free base amino acids. Therefore, L15 media would prevent extreme pH shift in ambient air during the collection. In addition to embryo benefits, such as protease inactivation, FBS also contains high levels of protein (i.e., albumin), preventing the egg/embryo from sticking to the bottom of the dish.
2. Turn on a dissecting/stereo microscope with a heat stage set to 37 °C.
3. Gather the following (Figure 2C):
   1. One 35-mm dish with a drop of 1 mL L15 + 1%FBS at 37 °C to wash blood off tissues.
   2. One 35-mm dish with 500 µL L15 + 1%FBS at 37 °C for embryo flushing (2 of these if collecting morula/blastocysts at 3.5 dpc).
   3. One 35-mm dish with 15 µL droplets of L15 + 1%FBS covered in 2–3 mL mineral oil at 37 °C.
      NOTE: If collecting zygotes at 0.5 dpc, an additional 35-mm dish containing two 100 µL drops of L15 + 1%FBS at 37 °C and one 100 µL drop of 0.1% hyaluronidase mixed in phosphate-buffered saline (PBS) or L15 + 1%FBS at 37 °C is needed.
   4. One 1-mL syringe with a blunted 30-G needle filled halfway with L15 + 1%FBS at 37 °C.
   5. Two glass capillary pipettes with different tip widths (Figure 2D). One larger for cumulus-oocyte-complexes, and one smaller for individual embryos.
   6. One glass capillary pipette with a rubber bulb attached, a mouth pipette tubing apparatus, or a homemade mouth pipette using tubing connected with a 0.22-µm filter and barrier P1000 tips, depending on preference (Figure 2D). Note that using a plastic pipette tip can result in embryos 'sticking' to plastic and becoming 'lost'.

4. Collecting the oviduct at 0.5 dpc

NOTE: At this time point, collections need to be completed by 11:00 h. If unavoidable, and the zygotes are collected later, embryos will need to be flushed using method 5 since the zygotes may be free of cumulus cells.

1. Euthanize mice per approved ACUC protocol.
2. Place the mouse in a supine position, spray the abdomen near the hind legs with 70% ethanol, and smooth the hair downward toward the feet.
3. Pick up the skin of the lower abdomen with forceps, make a small incision (~1–2 cm), hold the tail or legs of the mouse, and pull back the first skin layer to reveal the inner lining.
4. Pick up the abdominal muscle layer with forceps and make an incision in a V-shape, and flap the muscle over to expose the peritoneal cavity.
5. Push the intestine toward the head of the mouse and pull the abdominal fat down toward the feet to reveal the reproductive tract (Figure 3A). Ovaries will be covered with a gonadal fat pad and are visible near the kidneys, and the cervix will be near the bladder (Figure 3A).
6. Using the forceps, gently lift the uterus near the oviduct to gain clear visibility of the oviduct and the ovary.
   NOTE: The oviducts are very delicate and easily tear; do not grab the oviduct.
7. While the uterus is lifted, use a small pair of scissors to poke a hole underneath the oviduct (Figure 3B) and gently press down to separate the connective tissues and free the oviduct from the body cavity. Do not let go of the forceps holding the uterus.
8. Gently lift up the uterus and snip the area between the end of the oviduct and the ovary by placing scissors against the ovary (Figure 3C). This procedure will leave the ovary and ovarian fat pad behind inside the body cavity – the image in Figure 3D is shown outside the body cavity for clarity.
9. Cut again just below the utero-tubal junction (UTJ) connecting the oviduct to the uterus.
10. Place oviducts into 1 mL of L15 + 1%FBS media. The following procedures are performed under a light microscope.
11. Trim off fat if applicable.
12. Place the dissected oviducts into a clean 500 µL droplet of L15 + 1%FBS media.
13. Use scissors or the tip of the forceps to gently make a slit in the ampulla region of the oviduct. This region will be identified by eye; it will be swollen and translucent, with visible COCs clustered together.
    NOTE: If the media drop is bloody, move the tissues to the new drop before further dissection.
14. After making a small slit, use forceps and gently press on both sides of the bulging ampulla. The COC mass should slide/pop out into the media.
15. Using a glass capillary pipette with a larger tip width, connected to the mouth-pipette piece or a rubber bulb, move the COCs to 0.1% hyaluronidase in PBS or L15 + 1% FBS for approximately 15 s.
16. Watch the COC mass at all times; when the cumulus cells begin to fall away from the embryos, immediately move the zygotes to the adjacent 100 µL drop of L15 + 1%FBS to avoid further digestion from hyaluronidase, which could be harmful to the embryos.
17. Wash the embryos off of hyaluronidase one more time in the adjacent 100 µL drop of L15 + 1%FBS.
18. Move embryos into a droplet of one of the 15 µL drops of L15 + 1%FBS covered in mineral oil. At this point, switch to a glass capillary pipette with a smaller tip width.
19. Repeat steps 3 and 4 with the second COCs from the contralateral side of the oviduct. Once all embryos are in a 15 µL droplet of L15 + 1%FBS, move them through these droplets until no cumulus cells remain. Place the dish on a heated stage until imaging.
    NOTE: Zygotes can be identified by the presence of 2 pronuclei (Figure 4A). Eggs and embryos are encapsulated by the zona pellucida (ZP) (Figure 4A–C).

5. Collecting the oviduct at 1.5–2.5 dpc

1. Following steps 4.1–4.12, once oviducts are dissected and placed in L15 + 1%FBS and placed in 500 µL of L15 + 1%FBS, identify the infundibulum and its opening (ostium, Figure 5A). Look under the microscope for this entire procedure to ensure the correct placement of the needle.
2. Use the forceps to gently hold the infundibulum and insert the tip of a blunted-end 30-G needle into the ostium of the oviduct (Figure 5B). While inserting the needle, hold it as if holding a pencil.
3. Use forceps to gently glide the infundibulum up the needle shaft (similarly to pulling a sock up the foot). Do not forcefully push the needle into the infundibulum.
4. Once the needle is placed in the infundibulum, gently pin the oviduct to the bottom of the dish to secure its position and prevent it from sliding away during flushing.
5. Free the left hand holding the forceps and use the free left hand to push the media until the liquid and luminal content are visibly flushed out from the UTJ end of the oviduct. Do not look away during this process. The entire tube should puff up with liquid. This takes ~50–100 µL of media to flush through the tube.
6. Remove the flushed tissues from the dish.
7. Collect the embryos using a small-tip-width glass capillary pipette and place them in a 15 µL droplet of L15 + 1 % FBS medium.
   NOTE: Embryos will appear as perfect circles that sink to the bottom of the dish. The dish can be swirled lightly in a circular motion to center the embryos in the drop, but make sure to check the entire dish, especially around the edges, to verify that all embryos are retrieved.
8. Once all embryos are in a 15 µL drop of L15 + 1% FBS, move them through droplets until no other cell debris remains. Place the dish on a heated stage until imaging.
   NOTE: At 1.5 dpc and 2.5 dpc, embryos should be 2-cells and 4- to 8-cells, respectively (Figure 6).

6. Collecting the oviduct and uterus at 3.5 dpc

NOTE: By around 13:00 h on 3.5 dpc, the embryos will be in the uterus and should be at the morula (compacted embryos) or blastocyst stage (with a visible blastocoel or cavity formed). Rather than snipping the oviduct away from the uterus, the entire uterus will be removed.

1. Once the reproductive tract is visible, poke a hole underneath the oviduct to free it from the connective tissue. Then, at the cervix, lift the uterus, trim away connective tissue and fat, and trim upward toward the oviduct and ovary. Repeat on the contralateral side of the uterus.
2. Cut the oviduct away from the ovary (as indicated in Figure 3C) and the uterus near the cervix.
3. Place the reproductive tract in a dish with 1 mL of L15 + 1% FBS, and trim away any remaining fat and connective tissue if necessary.
4. Move the uterus with oviducts attached to a clean 500 µL L15 + 1% FBS medium droplet (Figure 7A). Under the stereo microscope, using the forceps to hold the upper uterine horn in place, insert the needle at the top of the uterus near the UTJ (Figure 7B) and flush the uterine lumen with ~500 µL/horn.
   NOTE: When flushing, the uterus is visibly inflated, and the flushed media flows through the cervical end into the dish. Repeat the flushing on the contralateral side of the uterine horn. Be careful not to push the liquid too hard, or the media may squirt out of the dish.
5. If needed, cut the oviduct near the UTJ from both sides of the uterus and place the dissected oviducts in the new clean drop of 500 µL L15 + 1%FBS and repeat the oviduct flushing as described above. Remove the uterus from the flushed dish.
6. Swirl the dish to bring the embryos to the center. Care should be taken when swirling the dish to avoid disturbing individual drops.
7. Collect all embryos from the uterus and oviduct (if needed) and place in a separate 15 µL drop of L15 + 1%FBS on a heated stage until imaging.
   NOTE: At 3.5 dpc, embryos should be at the morula or blastocyst stages (Figure 8). If there are no embryos in the uterus at 3.5 dpc, flushing the oviduct is required. It may indicate an embryo transport defect.

7. Imaging the collected embryos

1. When preparing for imaging, use the glass capillary tube to move the fluid around the embryos, pushing them toward the center of the droplets, and grouping them together as much as possible.
2. Use an inverted microscope with 10× and 20× objectives to image the embryos and determine their developmental stage. Color or monochromic imaging can both be used, depending on availability and preference.
   NOTE: While a dissecting/stereo microscope can be used to roughly identify embryonic stages, higher magnification is important for detecting detailed differences between groups.

Results

Vaginal smears were collected and analyzed following step 1. Representative results are shown in Figure 1B. The presence of nucleated epithelial cells, some cornified (enucleated cells), and a few leukocytes is categorized as a proestrus stage. At estrus, almost all epithelial cells are enucleated cornified cells (corn flake-like cells). During metestrus, leukocytes are present in large numbers surrounding the cornified cells. Leukocytes are much smaller, round cells compared to other cell t...

Discussion

Proper cycling can be evaluated using vaginal cytology. The average cycle in C57Bl/6J wild-type mice is 4.5 days^4,7. Collecting vaginal smears and documenting cell type can indicate whether the timing is correct and whether perturbations (disease state, drug treatment, gene deletion) could impact steroid hormone production. It is recommended that those new to evaluating vaginal smears perform hematoxylin and eosin staining to clearly differentiate cells and becom...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Alcohol lamp	Fisher Scientific	50-307-0254	70% Ethanol
B6D2F1/J mice	Jackson Laboratory	Stock #000664100006
Bulbs for small pipette	Fisher Scientific	03-448-21	1 mL to connect to glass capillary pipette
C57BL/6J mice	Jackson Laboratory	Stock #000664
Carbon dioxide	AirGas	CD USP50	Medical grade for euthanization
Diamond scribes	Fisher Scientific	08-675	Cutting glass pipettes
Ethanol	Decon Laboratories INC	2801	95%, 190 proof
Ethanol	Decon Laboratories INC	2701	100%, 200 proof
Fetal bovine serum	Sigma-Aldrich	12306C	Cell culture grade - heat inactivated
Forceps	Fisherbrand	16-100-110	Curved medium point general purpose forceps
Glass slides	Fisher Scientific	1255015	Positive charged for collecting vaginal cells
Hyaluronidase	Sigma-Aldrich	H4272	Type IV-S, powder, suitable for mouse embryo cell culture
Leibovitz’s L-15 medium	Gibco	41300-039	Powder, follow the manufacturer's instructions to make medium
Microscope for cytology	Leica Microsystems	DM1000LED	For mouse estrus staging
Microscope for embryo imaging	Leica Microsystems	DMi8 automated	Equiped with K3C camera
Microscope Stero	Leica Microsystems	Mz10f	With spotlight illumination LED3000 SLI
Mineral oil	Irvine Scientific	9305	Embryo culture grade
Mouth pipette tubing	Millopore Sigma	BR25931	BRAND spare adapter for micro-classic pipette controller
Needles	Fisher Scientific	14-826F	BD PrecisionGlide, 30G
Paper towel	Fisher Scientific	19120476	Scott multi-fold 4000/CS
Petri dishes	Falcon	351008	Size: 35 mm x 10 mm
Phosphate Buffered Saline (PBS)	Millipore Sigma	D8537	For making 0.1% hyaluronidase
Pipette tips - 200 µL	MidSci	PR-200RK-S	200-µL PR1MA pipette tips, sterile
Pipette tips - 1000 µL	MidSci	PR-1250RK-S	1250-µL PR1MA pipette tips, sterile
Pipetter 100 to 1000 µL	Eppendorf	2231300040	Eppendor Research Plus
Pipetter 20 to 200 µL	Eppendorf	2231300038	Eppendor Research Plus
Pipettes borosilicate glass pasteur	VWR	89399-924	Non-Plugged, 2 ml, 7 mm, 9", 5"
Saline	Pfizer Hospital	NDC 0409-4888-10	0.9% Sodium chloride, USP, sterile
Scissors dissecting (spring)	Roboz Surgical	RS-5610	For disecting oviduct
Scissors surgical	Fisher Scientific	12-460-788	For cutting skin and muscle tissue
Spray bottles 1 L	Fisherbrand	12-800-200	For holding 70% EtOH
Syringe	Fisher Scientific	14955464	1 mL, sterile
Syringe	BH Supplies	BH10LL	10 ml, sterile
Syringe filters	Foxx Life Sciences	3782415OEM	0.22-µm sterilizing membrane
Thermoplate	Tokai Hits	TPX series	For Leica TL bases
Toothpicks	Fisher Scientific	S24559	Wooden flat, for copulatory plug check
Tubing E-3603	Tygon	ACF00011	To connect with filter and 1250-µL pipette tip with barrier for homemade mouth pipette
Tweezers (Watch maker forceps)	Fisher Scientific	NC0410614	Roboz Surgical Dumont Tweezers; Pattern #5