Methods for Studying Uterine Contributions to Pregnancy Establishment in an Ovariectomized Mouse Model

Summary

Pregnancy establishment is a dynamic process involving complex embryo and uterine crosstalk. The precise contributions of the maternal uterine environment to these processes remain an active area of investigation. Here, detailed protocols are provided to aid in designing in vivo animal models to address these research questions.

Abstract

For pregnancy to be established, a viable blastocyst must successfully interact with a receptive uterine lining (endometrium) to facilitate implantation and placenta formation and enable ongoing pregnancy. The limitations to pregnancy success caused by embryonic defects are well known and have been largely overcome in recent decades with the rise of in vitro fertilization (IVF) and assisted reproductive technologies. As yet, however, the field has not overcome the limitations caused by an inadequately receptive endometrium, thus resulting in stagnating IVF success rates. Ovarian and endometrial functions are closely intertwined, as hormones produced by the ovary are responsible for the endometrium's menstrual cyclicity. As such, when using rodent models of pregnancy, it can be difficult to ascertain whether an observed result is due to an ovarian or uterine deficit. To overcome this, an ovariectomized mouse model was developed with embryo transfer or artificial decidualization to allow the study of uterine-specific contributions to pregnancy. This article will provide instructions on how to perform ovariectomy and offer insights into various techniques for supplying exogenous hormones to support successful artificial decidualization or pregnancy following embryo transfer from healthy donors. These techniques include subcutaneous injection, slow-release pellets, and osmotic mini pumps. The key advantages and disadvantages of each method will be discussed, enabling researchers to choose the best study design for their specific research question.

Introduction

With the rising use of assisted reproductive technologies in recent decades, many barriers to conception have been overcome, allowing many couples to start families despite fertility problems¹. Oocyte or sperm deficits can often be bypassed using in vitro fertilization or intracytoplasmic sperm injection; however, issues related to the uterus and endometrial receptivity remain an elusive "black box" of reproductive potential².

Pregnancy is established when a high-quality embryo successfully interacts with a receptive endometrium (uterine lining). The chances of successful pregnancy in any given menstrual cycle are low, at around 30%^3,4. Of those that are successful, only 50%-60% advance past 20 weeks of gestation, with implantation failure being responsible for 75% of pregnancies that do not reach 20 weeks³. Despite these figures dating back to the late 1990s, the field is yet to overcome the limitations caused by an inadequately receptive endometrium. This has resulted in stagnating - and sometimes declining - IVF success rates in recent years^5,6.

Women with unexplained infertility often have a displaced window of receptivity or are unable to achieve receptivity for unknown reasons. Recently, the endometrial receptivity array was developed, which assesses the expression of hundreds of genes with the purpose of tailoring the timing of embryo transfer to an individual's window of receptivity^7,8,9. However, the field still lacks an understanding of the pathogenesis of pregnancy complications that manifest after the implantation process is complete.

The female reproductive system is highly dynamic and under tight hormonal control. The hypothalamic-pituitary-gonadal (HPG) axis controls the release of luteinizing hormone and follicle-stimulating hormone, which regulate aspects of the ovarian cycle, including follicle maturation and estrogen and progesterone activity. In turn, the uterine menstrual cycle is regulated by estrogens and progesterone^10,11. Thus, studying uterine biological mechanisms is complicated by ovarian influence. For example, when studying how cancer therapies may impact the uterus, it can be difficult to distinguish if any uterine phenotype observed (such as pregnancy loss or menstrual acyclicity) is the result of a direct insult to the uterus or a consequential effect from damage to the ovaries.

To comprehensively understand fertility, the uterine contributions to pregnancy must be characterized. Importantly, this understanding must extend beyond uterine function under ovarian control. This cannot be studied in humans; therefore, animal models are often employed. As such, ovariectomy (OVX) is commonly used to enable researchers to regulate rodent estrous cycles (analogous to the menstrual cycle) by supplying hormones exogenously. Additionally, OVX allows uterine responses to be studied independently of ovarian influence¹². However, if hormones are not immediately supplied post-OVX, a menopause phenotype will eventuate, which needs to be carefully considered by the researchers.

OVX is frequently utilized in rodent models^{13,14,15,16,17} and is relatively easy to perform after adequate training. Methods vary depending on whether the ovary alone or the ovary and oviduct are removed, as well as depending on the age of the animal (adult, cycling animals have larger ovaries with a visible corpus luteum on their surface, meaning their ovaries are easier to visualize). Similarly, many methods of hormone supplementation exist, including subcutaneous injections¹⁴, slow-release pellets¹⁵, osmotic mini pumps¹⁸, and ovarian grafting.

In this article, detailed instructions are provided on how to perform ovariectomy and prepare three types of hormone supplementation, including subcutaneous injections, slow-release pellets, and osmotic mini pumps. Two detailed protocols are provided for experimental endpoints that benefit from OVX followed by exogenous hormone supplementation (embryo transfer and artificial decidualization). This article discusses the strengths and weaknesses of each approach with the goal of guiding researchers regarding how to perform studies to isolate the impacts on the uterus, specifically in the pregnancy and fertility fields of research.

Protocol

All animals were housed in temperature-controlled, high-barrier facilities (Monash University Animal Research Laboratory) with free food and water access and a 12 h light-dark cycle. All the procedures were performed in accordance with approval from the Monash Animal Research Platform Ethics committee (#21908, 17971) and performed in accordance with the National Health and Medical Research Council Code of Practice for the care and use of animals.

1. Surgical preparation

1. Autoclave all the surgical instruments, gauze, and paper towels required for the procedures on a hard/dry goods cycle at 121 °C with a hold time of 30 min and a drying time of 30 min.
2. Lay out a sterile bench pad for the surgical workspace and prepare analgesics.
   1. Dilute carprofen in sterile saline to 1 mg/ml, and dilute bupivacaine in sterile saline to a 0.5% (w/v) solution.
   2. Add 3.5 mL of meloxicam to a 400 mL cage water bottle.
3. Pre-warm the heat pad(s) for the recovery cages, and set up heat lamps to shine indirect light on the recovering animals.
4. Ensure all the appropriate PPE is worn, including a hair net, face mask, gown, and gloves.
5. Practice good sterile technique, including regularly spraying the gloves with ethanol and allowing them to evaporate before handling the animal or surgical tools to avoid ethanol contamination.

2. Performing ovariectomy

1. Using a gas anesthesia machine with isoflurane, prefill the induction box for 3-5 min at 5% isoflurane with the flow rate set at 4 L/min.
2. Place the mouse inside the induction box and, once unconscious, move it to the nose cone and reduce the flow rate to 0.4 L/min, with the isoflurane vaporizer set to ~2.5%.
   NOTE: The percentage of isoflurane used for the remainder of the procedure varies based on mouse strain, age, and exposure to treatments (e.g., chemotherapy) and should be adjusted based on a close assessment of each individual animal's breathing patterns. Breathing patterns should remain as regular abdominal breaths. Rapid, thoracic breaths may indicate that a deep surgical plane has not been reached or maintained; in this case, adjust the isoflurane vaporizer percentage as necessary.
3. Apply eye lubricant generously by squeezing the tube and gently dabbing the eye.
4. Shave a small (2 cm x 2 cm) area at and below the hunch of the spine.
5. Administer 5 mg/kg carprofen from a diluted solution of 1 mg/mL subcutaneously at the scruff of the neck.
6. Test the depth of anesthesia with the toe pinch reflex by pinching the mouse's back toe. If there is no toe pinch reaction, the animal is in the deep surgical plane, and the procedure can continue.
7. Apply betadine to the surgical area, and cover with a surgical drape (a gauze with a 2 cm x 2 cm window cut out).
8. Using rat-toothed forceps, pull the skin at the hunch of the back upward, and make a ~5 mm longitudinal incision.
   NOTE: A skin incision at this height on the animal's back is best for surgical clip placement to reduce the chance of the animal removing the clips and requiring clip repairs.
9. Using blunt forceps, proceed to blunt dissect the skin away from the underlying muscle layer, moving down and to one side toward the kidney.
10. Identify the kidney, ovary, and ovarian fat pad visually through the muscle wall.
    NOTE: The kidney will appear a dark red color, the fat pad will appear bright white, and if visible, the ovary will look like a small pink dot within the fat pad.
11. Using forceps, grab and lift the muscle layer. Make a ~0.5-1 cm incision with sharp surgical scissors. Continue holding the muscle wall with forceps, and change from scissors to blunt forceps to pull the ovarian fat pad through the incision.
12. Using a curved needle holder, clamp underneath the ovary and oviduct at the distal end of the uterine horn.
    NOTE: Alternatively, the ovary alone can be removed, leaving the oviduct intact. However, a dissecting microscope is required to accurately visualize the distinction between the ovary and oviduct.
13. Remove the ovary with scissors or a scalpel. Continue to clamp for 30 s to avoid excessive bleeding.
14. Remove the clamp, and dab it with sterile gauze if necessary.
15. To close the muscle wall incision, use forceps to lift the top of the incision so that the incision naturally pulls together.
16. Use silk sutures (size 3-0) to close the muscle wall incision with a surgeon's knot.
17. Apply two to three drops of bupivacaine topically using a 1 mL syringe without a needle attached, and repeat steps 2.9-2.17 on the other side.
18. To close the skin incision, use gauze to dab the area dry of excess bupivacaine, and press the two sides of the skin together.
19. Apply one to two 7 mm surgical clips, allowing space for swelling as part of the healing process.
20. Move the mouse to a recovery cage, and monitor closely for 15 min.
    NOTE: The animals should wake quickly; be sure to monitor breathing closely for normal thoracic breathing patterns.

3. Hormone preparation: Subcutaneous injection

1. Make a 1 mg/mL stock solution of estradiol.
   1. Weigh out 0.001 g (1 mg) of estradiol powder into a 1.5 mL sterile tube.
   2. Add 1 mL of 100% ethanol to the tube, and vortex for a few seconds.
      NOTE: The ethanol will remain clear with visible flecks of estradiol powder.
   3. Wrap the tube with film to prevent any ethanol evaporation.
   4. Wrap the tube in aluminum foil, and place it on a rocker overnight to completely dissolve the estradiol powder.
   5. Dilute this 1 mg/mL stock in sesame oil to the desired final concentration.
      NOTE: Doses of 100 ng for 3 days are required for priming prior to artificial decidualization, and additional low doses of 25 ng are required when progesterone is given. This is to combat the feedback loop controlling progesterone receptor expression. For embryo transfer, two doses of 100 ng on day 1 and day 3 prior to embryo transfer on day 4 are required. At the time of embryo transfer, a low dose of 25 ng is also required.
   6. Draw up the required amount of estradiol in oil into a 1 mL syringe, and then attach a 26 G needle tip.
   7. Inject the appropriate dose subcutaneously (either at the scruff or flank; 100 ng/100 µL or 25 ng/100 µL for priming prior to embryo transfer or artificial decidualization or at the time of embryo transfer) at the required frequency.
      NOTE: Oil is very viscous, so be sure to inject slowly and pause for a few seconds before removing the needle. This will minimize the amount of oil that leaks out of the injection site.
2. Make a 200 mg/mL stock solution of progesterone.
   1. Weigh out 0.4 g (400 mg) of progesterone powder into a 5 mL sterile tube.
   2. Add 2 mL of 100% ethanol to the tube, and vortex for a few seconds.
      NOTE: The ethanol will become white in color.
   3. Repeat steps 3.1.3-3.1.4.
   4. Dilute the 200 mg/mL stock in sesame oil to the desired final concentration.
      NOTE: Doses of 2 mg daily are required for supporting embryo transfer.
   5. Inject the appropriate dose (e.g., 2 mg/100 µL daily for supporting pregnancy) subcutaneously as in steps 3.1.6-3.1.7.

4. Hormone preparation: Slow-release pellets

1. Lay a foil across the surface of a laminar flow or class II biosafety hood.
2. Place all the equipment (gloves, Petri dishes, 1 mL syringes, fine forceps) in the hood, and switch on the UV for 20 min.
   NOTE: Do not switch on the UV with the sealant inside the hood as it will set.
3. Wash the silastic tubing in 100% ethanol, and allow it to air-dry in the hood. Once dry, mark ~1 cm lengths along the tubing, and cut with a scalpel.
4. Remove the plunger from a syringe, and squeeze in ~200 μL of sealant. Replace the plunger, and depress a small amount of sealant out of the syringe.
5. Apply a small amount of sealant to one end of the tubing, and smooth it over with a gloved finger.
6. Allow to dry overnight or for 20-30 min in UV light inside the hood.
7. Pour an appropriate amount of progesterone into a sterilized Petri dish. Using forceps, scoop pellets into the progesterone powder to fill the pellet.
   1. Tap the sealed end of the pellet on the hood surface to condense the progesterone down. Alternatively, use the end of sterilized forceps to stuff the progesterone down. Allow enough room for more sealant.
8. Seal the open end with sealant, as described in steps 3.4-3.5.
9. Wrap the Petri dish containing the progesterone pellets in foil to protect it from light.
10. Activate the pellets for a minimum of 72 h before subcutaneous insertion by incubating in 1% charcoal-stripped FCS (cs-FCS: PBS) at 37 °C.
    NOTE: Pellets can be made in bulk with a single sealed end in advance. However, fresh progesterone should be used to fill them each time. Ensure the pre-made pellets are UV-sterilized prior to filling with progesterone. The pellets will secrete ~500 µg/day for 6-10 days, which is sufficient support for artificial decidualization and embryo transfer procedures, though an additional low-dose estrogen injection may be required to maintain progesterone receptor activity beyond 4-5 days. Beyond 10 days, a replacement progesterone pellet may be required.

5. Hormone preparation: Osmotic mini pumps

1. Prepare progesterone at the desired concentration in an aqueous solution, and select the appropriate mini osmotic pump model (see Table of Materials).
   NOTE: For section 7 (experimental procedure: embryo transfer), the delivery of 2 mg/day for 12 days is required. Therefore, dissolve 28 mg of progesterone in ~100 µL of sterile water per animal (follow the manufacturer's instructions for the specific volume). Serial dilutions may be required. For section 8 (experimental procedure: artificial decidualization), the delivery of 500 µg per day for 3 days is required. Therefore, dissolve 1,500 µg of progesterone in ~100 µL of sterile water per animal. Prepare extra solution to account for the volume lost during the filling procedure.
2. Set up the equipment (gloves, low-lint wipes, Petri dishes, sterile saline, 1 mL syringes, small weigh boats, foil, and scales accurate to 0.01 g) within a class II biosafety hood, and then switch on the UV for 20 min.
3. Draw up the hormone solution into a 1 mL syringe, and then attach a sterile filling tube, carefully ensuring that there are no air bubbles.
4. Weigh the pump and its flow moderator within a sterile weigh boat.
5. Insert the filling tube through the opening at the top of the pump until it can go no further.
6. Holding the pump upright, slowly push the plunger of the syringe to fill the tube.
   NOTE: Rapid filling should be avoided as this can introduce air bubbles into the pump.
7. When the solution overflows from the top of the pump, gently remove the filling tube, and wipe off the excess solution with a sterile low-lint wipe.
8. Insert the flow moderator through the opening at the top of the pump until it can go no further. Once fully in, firmly press the pump and flow moderator together.
9. Weigh the filled pump with the flow moderator in place.
   NOTE: The difference in weight obtained from step 5.3 and step 5.8 will give the net weight of the solution loaded (i.e., a 0.1 g increase = 100 µL of solution added).
10. Place the filled pump into a sterile Petri dish filled with sterile saline.
11. Once all the pumps are filled, wrap the Petri dish in foil, and place it inside a 37 °C incubator to prime for at least 4-6 h (or until ready for use).

6. Surgical procedure: Insertion of subcutaneous hormone pellets and mini pumps

1. Prepare the area as per section 1 (surgical preparation).
2. Anesthetize the animals as per steps 2.1-2.3.
3. Shave a small area at the scruff of the neck (~1 cm x 1 cm).
4. Administer 5 mg/kg carprofen from a diluted solution of 1 mg/mL subcutaneously in the leg flank.
5. Test for the toe pinch reflex. If there is no reflex, the animal is in the deep surgical plane, and the procedure can begin.
6. Apply betadine to the surgical area, and cover it with a surgical drape (a gauze with a 2 cm x 2 cm window cut out).
7. Using rat-toothed forceps, pull the skin at the scruff of the neck (halfway between the true scruff and the hunch of the back) upward, and make a ~5 mm longitudinal incision.
8. Using blunt forceps, blunt dissect the skin away from the underlying muscle layer in a downward direction.
   NOTE: For osmotic mini pump insertion, create a pocket along one side of the animal so that the pump does not restrict the animal's movement or press up against the incision site.
9. Once sufficient space has been made for the hormone pellet or mini pump, use sterile forceps to pick up the pellet or mini pump, and insert it into the subcutaneous pocket made with blunt dissection.
10. To close the skin incision, ensure the pellet or mini pump is far enough into the pocket that surgical clips will not damage it.
11. Topically apply bupivacaine as per step 2.17.
12. Close the wound with one surgical clip. Move the mouse to a recovery cage, and monitor closely for 15 min. As this is a short procedure, the animals should be ambulatory within minutes.

7. Experimental procedure: Embryo transfer

1. For ovariectomized animals, hormone-prime 3 days prior to embryo transfer by a subcutaneous injection of 100 ng/100 µL estradiol (step 3.1).
2. One day prior to the embryo transfer, hormone-prime the animals with a subcutaneous injection of 2 mg/100 µL medroxyprogesterone acetate (step 3.2).
3. Prepare the area as per section 1 (surgical preparation).
4. Anesthetize the animals as per steps 2.1-2.3.
5. Begin the procedure as per steps 2.4-2.10.
6. Under a dissecting microscope, create an intrauterine injection point with a 26 G needle tip.
7. Pipette five blastocysts into an M2 media drop, and then transfer these into the uterine horn.
8. To close the muscle wall incision, lift the top of the incision using forceps so that the incision naturally pulls together.
9. Using silk sutures, close the muscle wall site with a surgeon's knot. Apply drops of bupivacaine topically.
10. Repeat steps 7.5-7.8 on the other side.
11. To close the skin incision, use gauze to dab the area dry of excess bupivacaine, and press the two sides of the skin together.
12. Apply one to two surgical clips, allowing space for swelling as part of the healing process.
    NOTE: If the animals are ovariectomized, exogenous hormones are required at the time of embryo transfer. Either subcutaneously inject progesterone (2 mg) or insert a subcutaneous progesterone pellet or osmotic mini pump. To combat excess progesterone, a subcutaneous injection of low dose (25 ng/100 µL) estrogen is required at the time of embryo transfer.
13. Carefully carry the mouse to a recovery cage, and monitor closely for 15 min.
    NOTE: The animals should wake quickly; be sure to monitor breathing closely for normal thoracic breathing patterns.

8. Experimental procedure: Artificial decidualization

1. Hormone-prime the animals with 100 ng/100 µL estradiol on day 1, day 2, and day 3, as per section 3 (hormone preparation: subcutaneous injection) 8 days prior to artificial decidualization.
2. Hormone-prime the animals with 5 ng/100 µL estradiol on day 7, day 8, and day 9, as per section 3 (hormone preparation: subcutaneous injection) 2 days prior to artificial decidualization.
   NOTE: The final injection must occur a minimum of 3 h (and a maximum of 4 h) before the artificial decidualization procedure.
3. Hormone-prime animals with a subcutaneous progesterone pellet or mini osmotic pump (500 µg/day), as per section 4, section 5, and section 8, 2 days prior to artificial decidualization.
4. Prepare the area as per section 1 (surgical preparation).
5. Anesthetize the animals as per steps 2.1-2.2.
6. Test the depth of anesthesia with the toe pinch reflex. If there is no toe pinch reaction, the animal is in the deep surgical plane, and the procedure can continue.
7. Place the mouse in a prone position, lift the tail, and slowly insert a 6 mm diameter speculum into the vagina.
8. Whilst keeping the nose of the animal in the anesthetic nose cone, place the lower body of the animal between the first and second fingers of the non-dominant hand. Use the thumb to gently push the tail upward to keep the vaginal opening in view.
9. Transfer 20 µL of sesame oil into one uterine horn using a non-surgical embryo transfer tip (attached to a 20 µL pipette).
   1. Keeping the pipette level with the vaginal opening, insert the tip into the vagina and through the cervix into the uterine horn. Once the tip is in the uterine horn, gently press the tip against the endometrial surface (if using the handling technique above, this movement will be felt against the second finger), and slowly expel the oil.
      NOTE: Keep the pipette plunger depressed, wait for 10 s to ensure all the oil has dispersed, and slowly remove the transfer tip whilst keeping the plunger depressed.
10. Remove the speculum from the vagina.
11. Carefully carry the mouse to the recovery cage, and monitor closely for 15 min.
    NOTE: The animals should wake quickly. Monitor their breathing closely for normal thoracic breathing patterns.
12. Limit the handling of the animals, and keep them in a quiet environment for 96 h after the procedure.
    NOTE: Loud noises or abrupt changes to their light and dark cycle will impact the success of the procedure. At the time of tissue dissection, the extent of decidualization success can be measured as a ratio of uterine weight to body weight. The artificial decidualization procedure has an 80% success rate. Therefore, exclude animals that fail to decidualize, and factor this in when selecting sample sizes for the experiments.

9. Surgical procedure: Post-surgical recovery, monitoring, and clip repairs

1. Allow the animals to recover half-on, half-off heat pads overnight before returning them to their home cage.
2. Monitor the animals daily for 5 days post-surgery, paying close attention to the wound site for signs of infection.
3. Perform clip repair if necessary.
   1. Prepare the area as per section 1 (surgical preparation).
   2. Anesthetize the animals as per steps 2.1-2.3.
   3. Remove the existing clip using a clip remover if the clip is still present.
   4. Apply a new surgical clip, as per steps 2.19-2.21.
4. Remove the surgical clips 7 days post-surgery as per steps 9.3.1-9.3.3 or when the animal is next under anesthesia (such as for an embryo transfer, the artificial decidualization procedure, or a subcutaneous injection following surgery).

Representative Results

A well-characterized model of artificial decidualization is described in this protocol paper (Figure 1A). Here, young adult female mice (8 weeks old) underwent surgical ovariectomy as described in section 1 and section 2. The mice were then rested for 2 weeks to ensure that the endogenous ovarian hormones dissipated before being supported with exogenous hormones as described in sections 3-7 and section 9. Artificial decidualization was induced by an intravaginal injection of sesame oil, and then the animals were rested until tissue collection, as described in section 9. In this study, artificial decidualization was performed in C57BL6/J mice, a commonly used mouse strain. At the time of tissue collection, the body weight was recorded, and the uterus was dissected and well-trimmed before being weighed (Figure 1B). The extent of the decidual response was recorded by expressing the uterine weight as a ratio of the body weight. In this study, 80% of the C57BL6/J mice decidualized (0.01012 ± 0.001515, n = 15), while 20% of the animal uteri did not decidualize (0.002108 ± 0.0001764, n = 3) (Figure 1C).

Figure 1: Schematic and representative results. (A) Schematic timeline for experimentally inducing artificial decidualization in a mouse model. Abbreviations: OVX = ovariectomy; E2 = estradiol (100 ng days 1-3, 5 ng days 7-9); P4 = progesterone. Note: A P4 pellet was used to generate the results presented here. Alternative methods for progesterone delivery include daily subcutaneous injections and mini-osmotic pumps. (B) Representative images of non-decidualised (ND) and decidualised (D) uteri from young adult C57BL6/J mice. Scale bars = 5 mm.(C) Comparison of the uterine weight to body weight (UW:BW) ratio in non-decidualized and decidualized animals. Data are mean ± SEM; Mann-Whitney test, **p = 0.003; ND: n = 3, D: n = 15. Please click here to view a larger version of this figure.

Hormone delivery method	Strengths	Weaknesses
Subcutaneous injections	No surgical intervention required	Repeated daily handling
	Accessible technique that does not require surgical training (as compared to pellet implantation)	Hormones in oil can leak out of the injection site, therefore amount absorbed by each animal can vary
Slow release pellets	No need for daily handling	Surgical procedure required
	Can be made in house	Not commercially available
	Affordable alternative to osmotic mini pumps
	Small and very well tolerated by animals
Osmotic mini pumps	No need for daily handling	Surgical procedure required
	Commercially available	Expensive
	Most accurate delivery method	Much larger than slow release pellets

Table 1: Strengths and weaknesses of the hormone delivery methods.

Discussion

This article provides step-by-step instructions on how to perform OVX and provide exogenous hormones for studies focused on understanding the uterine contributions to pregnancy and fertility. Two detailed protocols are provided on two experimental applications of these methods, including performing embryo transfer and inducing decidualization artificially.

Whilst performing OVX can be challenging initially - especially for researchers new to rodent models - it is a relatively simple procedure once appropriately trained and practiced. The key steps in the procedures include closely monitoring the animals while they are under anesthesia and ensuring there is no ovarian tissue left behind. In some models, the oviduct may be left intact. However, it should be noted that the oviduct is a hormone-responsive tissue with abundant estrogen and progesterone receptors¹⁹. The surgical protocol for removing the ovary and oviduct is much simpler compared to removing just the ovary, as the former can be completed with the naked eye. To remove just the ovary and leave the oviduct in place in the latter case, a dissecting microscope is required, as this is a much more intricate procedure. Consequently, the operating time may be extended, as the animal needs to be moved between the dissecting microscope stage and the operating field for different parts of the procedure, such as suturing the internal body wall.

The analgesic protocols detailed here are standard and approved by the Monash University Animal Ethics committee, so they may vary depending on the individual institution's ethics committee requirements or preferences. It should be noted that no analgesia was provided for the artificial decidualization procedure, as typical non-steroidal anti-inflammatories interfere with the decidualization process. If researchers wish to provide analgesics at the time of artificial decidualization, this should be taken into consideration.

This work presents three methods of hormone delivery to supplement ovarian hormones following OVX, and each method has its own strengths and weaknesses (Table 1). Subcutaneous injections of hormones in oil are common in the literature^14,16,17. This technique has many strengths, including the fact that no surgical procedure is required, and, thus, no formal training in rodent surgery or gas anesthesia is required. This makes subcutaneous injection an accessible option for almost all research groups. Injections are also affordable and easy to carry out. Practically, however, they have some limitations, particularly in models of pregnancy. To maintain pregnancy in an OVX animal, hormone supplementation with progesterone must be given daily to support the pregnancy. It may be possible to stop the daily injections once the placenta is sufficiently developed to take over as the main source of progesterone, though this has not been trialed in the protocols presented here. Anecdotally, it is possible for hormones in oil to leak from the injection site following subcutaneous injection. In part, this may be due to the needle size required (26 G) to easily dispense something as viscous as sesame oil. Therefore, this leakage needs to be monitored and recorded when performing injections in oil in order to correlate this with the experimental outcomes.

Slow-release pellets are preferable to subcutaneous injections, as they are cost-effective and simple to make in-house. However, they require multiple overnight steps, which should be considered when planning experimental timelines. These pellets secrete approximately 500 µg daily (as assessed during a time course incubation in cell culture medium and subsequent progesterone ELISA). It should be noted this is a lower concentration compared to the daily subcutaneous injections described above, and this is due to the consistency in the delivery of progesterone from the pellet. As aforementioned, oil injections can leak out of the injection site, thus reducing the overall concentration delivered. In previous studies, these pellets have only been active in vivo for up to 10 days after they are surgically inserted. Therefore, in pregnancy studies, it remains unclear if it may be necessary to insert a second pellet at mid-gestation or whether the placenta could sufficiently provide endocrine support for the pregnancy by that stage. These pellets are, therefore, optimal for short-term models of pregnancy, including the artificial decidualization protocol presented here, as well as pregnancy studies up to 10 days post-embryo transfer. While slow-release pellets negate the need for daily animal handling and injections, some low-dose estrogen injections are still required to balance the progesterone receptor feedback loop. This strategy has been used previously^20,21.

Lastly, osmotic mini pumps are the most accurate hormone delivery method and are commercially available, but they are the most expensive option. Osmotic mini pumps can deliver a set concentration of hormone daily for up to 28 days, depending on the model selected. Similar to the slow-release pellets, while osmotic mini pumps avoid the need for daily animal handling, some low-dose estrogen injections are still required.

The artificial decidualization protocol described here allows the study of an early pregnancy milestone independent of ovarian and embryonic influence. While humans undergo decidualization with every menstrual cycle, rodents only decidualize during pregnancy establishment. Thus, this model has immense value for studying human-like pregnancy milestones in a manipulable rodent model. The procedure detailed herein is relatively non-invasive, as it uses a non-surgical embryo transfer device (NSET) to deliver sesame oil directly to the uterine horn via the vagina and cervix. Though this procedure is less invasive than other methodologies, it can become quite expensive when using commercial NSETs. In comparison, other published models of artificial decidualization require a surgical procedure to perform intrauterine injections of oil¹⁷. This requires a surgical setup similar to that described in section 1 and steps 2.1-2.11. However, in animals that have been ovariectomized previously, it can be more challenging to identify and expose the uterine horn. There can also be adhesions formed from the previous surgical procedure for ovariectomy. Thus, while it may be more cost-efficient to perform surgical intrauterine injections to induce decidualization, the surgical and anesthesia time is substantially longer than the alternative of using NSETs. There are established protocols for in-house fabricated alternatives²² to commercially available NSETs, which are much more cost-effective.

While the embryo transfer procedure is described here, we have previously published this model and its success rates across different strains of mice¹⁴. Moreover, while the embryo transfer method described here uses a surgical approach, NSETs could also be integrated into this procedure.

Future directions in this area should include studies focused on the specific uterine contributions to the establishment and maintenance of pregnancy. This knowledge is critical for furthering our understanding of idiopathic infertility, implantation failure, and pregnancy complications. Expanding our knowledge in these areas is also fundamental for improving clinical outcomes for IVF/ICSI patients, as well as our comprehension of pregnancy as a biological process.

In conclusion, OVX is a simple procedure that can be integrated into animal models to study the uterine contributions to pregnancy and fertility. Models in the future will benefit from integrating OVX and exogenous hormone delivery so that comparisons can be made between ovarian-specific and uterine-specific contributions to fertility and pregnancy.

Materials

Name	Company	Catalog Number	Comments
ALZET 1002 mini osmotic pumps	BioScientific	1002	Delivers 0.25 µL/h for 14 days. Use for section 7 (Experimental procedure - Embryo transfer).
ALZET 1003D mini osmotic pumps	BioScientific	1003D	Delivers 1 µL/h for 14 days. Use for section 8 (Experimental procedure - Artificial decidualization).
ALZET Reflex 7 mm clips	BioScientific	0009971	Either Reflex clips or Michel clips can be used for wound closure, depending on preference
ALZET Reflex clip applicator	BioScientific	0009974	Either Reflex clips or Michel clips can be used for wound closure, depending on preference
ALZET Reflex clip remover	BioScientific	0009976	Either Reflex clips or Michel clips can be used for wound closure, depending on preference
Bupivicaine injection	Pfizer	NA	Stock 0.5%. Use at 0.05% in saline
Estradiol	Sigma	E8875
Meloxicam	Ilium	NA	Active constituent 0.5 mg/mL. Use 3.5 mL per 400 mL cage water bottle, or as your institution's vet prescribes.
Michel clips	Daniels	NS-000242
Multi purpose sealant	Dow Corning	732
Non-surgical embryo transfer (NSET) device	ParaTechs	60010	Contains 6 mm speculum. Single use only.
Progesterone	Sigma	P0130	Soluble in ethanol. Use for  section 3 (Hormone preparation - subcutaneous injection) and  section 4 (Hormone preparation - slow-release pellets)
Progesterone	Sigma	P7556	Soluble in water. Use for section 5 (Hormone preparation - osmotic mini pumps)
Refresh eye ointment	Allergan	NA	42.5% w/v liquid paraffin, 57.3% w/v soft white paraffin
Rimadyl Carprofen	Zoetis	NA	Stock 50 mg/mL. Use at 1 mg/ml (for 5 mg/kg dose)
Rubber tubing	Dow Corning	508-008	Washed in 100% ethanol and cut into 1 cm pieces. Inside diameter 1.57 mm ±  0.23 mm; outside diamater 3.18 mm ± 0.23 mm; wall 0.81 mm.
Sesame oil	Sigma	S3547
Sofsilk Silk sutures size 3-0	Covidien	GS-832