Mouse surgical model of mechanical uterine injury and subsequent embryo defects

Abstract

Uterine injury from procedures such as Cesarean sections (C-sections) often have severe consequences on subsequent pregnancies, leading to disorders such as uterine placenta previa, placenta accreta spectrum (PAS), and Cesarean scar pregnancy. With rates of C-section at approximately 30% of deliveries in the US and projected to continue to climb, an understanding of the mechanisms by which these pregnancy disorders arise and opportunities for intervention are sorely needed. However, there are currently very few animal models of uterine injury and its subsequent impacts on maternal as well as in utero and postnatal fetal outcomes. Here we describe a procedure for a novel model of surgically-induced uterine injury in the genetically tractable laboratory mouse (Mus musculus). We describe preparatory steps for surgery, the induction of uterine injury itself, and post-surgical recovery. We then provide supporting information regarding downstream dissection of pregnant mice. Lastly, we include additional information regarding estrous cycle staging in order to perform surgeries and dissections at the relevant phase in non-pregnant mice.

BASIC PROTOCOL 1: Preparation for surgery

BASIC PROTOCOL 2: Surgery and induction of uterine injury

BASIC PROTOCOL 3: Mating and dissection of pregnant mice as endpoint analyses

SUPPORT PROTOCOL 1: Estrous staging of animals

INTRODUCTION:

Uterine injury resulting from procedures such as Cesarean sections (C-sections), myomectomies, and dilatation & curettage (D&C) often have severe consequences on later pregnancies, leading to disorders such as placenta previa, placenta accreta spectrum (PAS), implantation failure, and Cesarean scar pregnancy (Ang, Skokan, & McKinley, 2023; Chazotte & Cohen, 1990; Fitzpatrick et al., 2012; Jauniaux, Gronbeck, Bunce, Langhoff-Roos, & Collins, 2019; Miller, Chollet, & Goodwin, 1997; Wortman & Alexander, 2013). Although C-sections represent ~30% of deliveries in the US (Bowman et al., 2014; Miller et al., 1997; Solheim et al., 2011), an understanding of the mechanisms by which these pregnancy disorders arise is limited (Biswas, Sawhney, Dass, Saran, & Vasishta, 1999; Cramer & Heller, 2016; Khong, Cramer, & Heller, 2018; Khong & Robertson, 1987; Silver & Barbour, 2015; Stanek & Drummond, 2007; Tantbirojn, Crum, & Parast, 2008). Gaining insights in these questions, as well as developing a platform for testing new strategies for diagnosis and intervention, requires the use of animal models of uterine injury, which until recently have not been available (Buhimschi, Zhao, Sora, Madri, & Buhimschi, 2010; Burke, Zsengeller, Karumanchi, & Shainker, 2020; Li et al., 2020).

In this article, we thus describe a novel methodology for introducing uterine injury into mice. We then provide an example of subsequent downstream assessment of injury-related outcomes. Specifically, in protocol 1 we provide instructions for pre-operative procedures. In protocol 2, we detail the surgical procedure itself for the induction of injury to the uterine horn. Finally, in protocol 3, we describe dissection of a pregnant dam in order to assess the effects of uterine surgery on a subsequent pregnancy. In addition, in optional Support Protocol 1, we outline steps for estrous staging of animals in order to assess the impact of the estrous cycle on surgical outcomes.

STRATEGIC PLANNING

Since the estrous cycle greatly affects the state of the uterus in terms of cell type composition (Bertolin & Murphy, 2014), transcriptome (Roberson et al., 2022), and response to injury (Zhang et al., 2024), it is important to consider estrous phase in experimental design. Therefore, if a particular procedure such as surgery or dissection must be constrained to a particular day, advance planning is necessary to obtain enough animals such that they are at the desired phase of estrous by the day of the procedure. Alternatively, or in addition, the researcher may track the estrous cycle of animals intended for upcoming procedures and proceed when the animal is at the desired phase.

With respect to the procedure itself, an important consideration is to obtain the necessary approvals in advance for working with controlled substances such as ketamine and xylazine.

BASIC PROTOCOL 1: Preparation for surgery

Introductory paragraph:

In this protocol, we describe the steps required for preparing the animal for surgery. First, we provide information for preparing the surgery area in a procedure room. Second, we describe the steps for anesthetizing an animal using ketamine and xylazine. Lastly, we outline creating a sterile field in anticipation of the surgery itself.

All surgical procedures must obtain institutional animal ethics committee approval prior to use. Each institution may have different requirements with regards to anesthesia, surgical suites, and aseptic surgical techniques. Researchers must follow the guidelines as specified at and approved by their home institution.

It is important to achieve a secure scruff of the animal prior to administering any fluids intraperitoneally (I.P.) or subcutaneously (S.C.) for the safety of both the animal and researcher. Achieving a plane of anesthesia may be challenging, particularly when using ketamine/xylazine. Isoflurane supplied via a nosecone is an alternative for this procedure. Courses in animal handling, animal husbandry, and aseptic rodent surgery are recommended prior to attempting this protocol.

Materials:

Solutions and reagents:

Ketamine/xylazine solution (see Recipes)

• Ketamine HCl (e.g. Dechra, 100 mg/mL, 10 mL)

• Xylazine HCl (e.g. AnaSed, 100 mg/mL, 50 mL)

Buprenorphine SR or ER (ZooPharm, 0.5 mg/mL at 5 mL per vial)

Sterile ophthalmic ointment (Puralube vet ointment, SKU: PH-PURALUBE-VET)

Surgical tools:

Sterilization pouches (e.g. Fisher Scientific, cat. no. 19-910-673)

Needle holder (e.g. Fine Science Tools, cat. no. 12500-12)

Forceps (e.g. Fine Science Tools, cat. no. 11255-20)

Curved Iris scissors (e.g. World Precision Instrument, cat. no. 501759G)

AutoClip wound closing system (e.g. Fine Science Tools, 12020-00)

Wound clip refills (e.g. Fine Science Tools, cat. no. 12022-09)

Personal protective equipment (PPE):

Lab coat (e.g. Fisher Scientific, cat. no. 19-472-472)

Surgical hair cap (e.g. Uline, cat. no. S-10480BLU)

Surgical masks (e.g. Fisher Scientific, cat. no. 18-048-010)

Nitrile gloves (e.g. Fisher Scientific, cat. no. 19-130-1597C)

Additional surgical items and supplies for surgical and recovery areas:

Absorbent bench pads (e.g. Fisher Scientific, cat. no. NC9110121)

Animal cages (e.g. VWR cat. no. 10712-110 and 10715-323)

Scale (e.g. Fisher Scientific, cat. no. S93805)

Plastic beaker (e.g. Thermo Scientific, cat. no. 12010250)

BD Lo-Dose U-100 insulin syringe (Fisher Scientific, cat. no. 14-826-79)

Fur trimmer (e.g. Fisher Scientific, cat. no. 50-195-4544)

Lint roller (e.g. Fisher Scientific, cat. no. 17-018-329)

Sterile swabs for eye ointment application (e.g. Fisher Scientific, cat. no. 18-366-472)

Surgical drape (e.g. Fisher Scientific, cat. no. 50-209-1792)

Glad Press ‘n Seal (e.g. Fisher Scientific, cat. no. NC1089054)

Illuminator for surgeries (e.g. AmScope, cat. no. HL150-AY)

Spray bottle (e.g. Fisher Scientific, cat. no. S413505P)

70% Ethanol (e.g. Fisher Scientific, cat. no. BP82031GAL)

Betadine swabsticks (e.g. Medline, cat. no. MDS093901)

Sterile alcohol prep pads (e.g. Fisher Scientific, cat. no. 22-363-750)

Warming/induction chamber with full floor warming (e.g. Fisher Scientific, cat. no. 14-370-302)

Protocol steps:

1. Prior to day of surgery, autoclave surgical instruments in a sterilization pouch.

2. At the time of surgery, don appropriate personal protective equipment (PPE): lab coat, hair cap, surgical facemask, and gloves.

3. Optional: To first identify animals in the desired phase of the estrous cycle, perform vaginal lavage followed by crystal violet staining and imaging (see Support Protocol 1).

4. Prepare the pre-operative area. This should consist of one absorbent bench pad lined with additional paper towels, flanked by one animal cage containing paper towels in which the animal may rest after receiving anesthesia prior to surgery.

5. Select the appropriate animal for surgery. Weigh the animal in a tared plastic beaker or similar container on a scale. Record the animal’s weight to calculate required amounts of anesthetic, analgesic, and saline. See Figure 1A.

6. Calculate the amount of anesthetic required to achieve a final dose of 100 mg/kg ketamine and 10 mg/kg xylazine. Fill an insulin syringe with the appropriate volume of a working solution of 10 mg/mL ketamine and 1 mg/mL xylazine diluted in 0.9% sodium chloride saline solution (e.g. 0.2 mL of ketamine/xylazine solution for a 20 g animal). Scruff and invert the animal. Inject the dose intraperitoneally (I.P.). Place the animal to rest in the pre-operative cage. Monitor animal and deliver an extra 0.02 mL of ketamine/xylazine if no cessation of motion is observed within ~3 minutes. Check for front toe pinch reflex after ~10 minutes. Animals typically reach a deep plane of anesthesia within 15 minutes. See Figure 1B and 1C.

   The amount of anesthetic required differs by strain. For example, C3H mice typically require approximately 20% more anesthetic than C57BL/6 or BALB/c animals (e.g. 0.24 mL of ketamine/xylazine solution for a 20 g animal). Dosing may need to be determined empirically based on each individual strain.

   To minimize injury to internal organs during I.P. injection, tip the animal back such that the head is lower than the abdomen. Insert the needle slightly above the lowest nipple.

   The needle should penetrate both the outer skin and the inner muscle wall. These are typically two different sensations of resistance upon needle insertion. If the needle is not inserted into the peritoneal cavity, a small subcutaneous bubble of liquid will form.

   Isoflurane delivered via nosecone is an acceptable alternative to ketamine/xylazine.

7. Inject the animal with 0.5 mg/kg Buprenorphine SR once the animal’s movements cease, generally within ~3 minutes of delivering ketamine/xylazine. See Figure 1D.

   Since the Buprenorphine SR both a very small volume and quite viscous, we suggest injecting near a shoulder blade of the animal while tenting the skin with the other hand.

8. Apply ophthalmic ointment to the eyes using a sterile cotton swab. See Figure 2A.

9. Place animal onto pre-operative absorbent pad with paper towels and shave the incision site at the lower-to-mid back See Figure 2B. Remove excess fur with paper towels, followed by a lint roller to remove remaining hairs. See Figure 2C.

   Care should be exercised during shaving to prevent cuts to the skin.

10. Disinfect incision site using alternating alcohol prep pads and betadine swabsticks, starting from the center of the incision site and wiping in a circular motion outward. Repeat for a total of 3 times. See Figure 2D and 2E.

11. Prepare a second absorbent pad under an illuminator for the surgery itself. Place a sterile drape over the absorbent pad. Spray gloves and illuminator with 70% ethanol to disinfect. Place autoclaved or sterilized surgical tools at the edges of the sterile surgical drape and animal in the center of the surgical pad. See Figure 3A.

12. Prepare a sterile field by placing a sheet of Glad brand Press ‘n Seal over the animal, leaving the snout exposed. Cut a small window over the shaved incision site. Rest surgical tools on a sterile surface when necessary during use. See Figure 3B, 3C, and 3D.

    Surgical drapes are an acceptable but costlier alternative to Press ‘n Seal. Press ‘n Seal is food-grade and thus considered sterile.

Figure 1:

Anesthetization of the animal subject.

A. Weigh the animal in a tared container set on a scale.

B. Using an insulin syringe, extract the appropriate volume of anesthetic required to achieve the desired final amount (mg/kg) to be administered to the animal.

C. Inject anesthetic (ketamine and xylazine) intraperitoneally into the animal by inserting the needle near the lowest nipples.

D. Inject analgesic (Buprenorphine) subcutaneously into the animal near a shoulder blade.

Figure 2:

Preparation of the animal subject for surgery.

A. Apply ophthalmic ointment to the eyes using a sterile cotton swab.

B. Shave the incision site.

C. After removing excess fur, remove any remaining hairs with a lint roller.

D. Swab the incision site with an alcohol prep pad.

E. Swab the incision site with a betadine swabstick. Repeat with alternating alcohol prep pads and betadine swabsticks for a total of 3 rounds.

Figure 3:

Preparation of the surgical space.

A. Place all sterilized instruments on a sterile drape. Position the anesthetized animal below the illuminator.

B. Position a piece of Press ‘n Seal such that the animal’s snout is exposed to air.

C. Cut a small opening in the Press ‘n Seal directly over the incision site.

D. View of the exposed incision site through the Press ‘n Seal opening.

BASIC PROTOCOL 2: Surgery and induction of uterine injury

Introductory paragraph:

In this protocol, we describe the surgical procedure for introducing uterine injury into a single uterine horn of a non-pregnant mouse. This procedure entails performing a dorsal laparotomy to expose the top of the uterine horn, followed by the insertion of a burred 25G needle into the uterine lumen and introducing a series of scrapes to damage the endometrium and/or to cut all the way through the uterine wall as well. At the end of the procedure, a longitudinal wound will have been created in the experimental uterine horn, allowing the researcher to address their question of interest, such as the molecular and structural trajectories of wound healing and how the injury introduced impacts a subsequent pregnancy. Care should be exercised when handling the needle to avoid injury to the researcher.

It is critical for the animal to be in a deep plane of anesthesia before beginning this protocol. As described in protocol 1, several parameters indicate that an animal is appropriately anesthetized – the cessation of movement, a reduction in respiratory rate, and most reliably, the absence of a front paw reflex. The rear paw reflex upon a light touch should also be absent; however, it is often the case, even at the appropriate plane of anesthesia, that this reflex is still present if strong pressure is applied.

Materials:

0.9% sodium chloride for injection, sterile (e.g. Farris labs, SKU: 63323-186-01)

Absorbable 4–0 vicryl suture (e.g. Ethicon, cat. no. J214)

25G needle (e.g. BD, cat. no. 305122) with tip bent or burred by pressing at a 45-degree angle against a sterile hard surface

Animal ear-punch (e.g. Fisher Scientific, cat. no. 13-812-201)

Carprofen (Zoetis, Rimadyl, 50 mg/mL at 20 mL per vial)

Dry glass bead surgical tool sterilizer (e.g. Fisher Scientific, cat. no. 10-000-632)

Protocol steps:

1. Using forceps and a pair of curved or straight Iris scissors, cut a ~2–3 mm incision in the mouse dorsal skin approximately 1/3 of the distance between the base of the tail and neck and approximately 0.5 cm from the midline. See Protocol 1 for catalog information. See Figure 4A.

2. Visually identify the ovarian fat pad under the fascia beneath the skin. Cut a ~2–3 mm incision in the fascia, parallel to the skin incision. See Figure 4B.

   The ovarian fat pad is a white mass that is located below the spleen.

3. Using a pair of forceps, gently grip the ovarian fat pad and begin extricating it from the incision in the fascia and the skin. See Figure 4C and 4D.

   It may be necessary to cut parts of the fat pad so that the ovary and uterus can be removed from the body cavity. Care should be taken to avoid injuring any part of the female reproductive tract itself, as well as to avoid the spleen or any large blood vessels near the fat pad.

4. Once the upper third of the uterus is exposed (Figure 4D), grip the uterotubal ligation with the forceps (Figure 5A). Create a small hole in the top of the uterus using the sharp end of the bent needle (Figure 5B). Insert the needle and begin drawing the needle out to scrape the antimesometrial surface of the uterine horn (Figure 5C). Continue scraping until the desired degree of damage is inflicted, including complete perforation of both the endometrial and myometrial layers. See Figure 5D.

   Care should be exercised to avoid ripping the uterus at the uterotubal ligation, which would result in separation of the uterine horn from the oviduct and ovary. This scenario can occur if excessive pressure is applied in inserting the needle into the uterus. If the researcher finds it difficult to create a small hole with the bent needle, an unbent 25G or 27G needle may be used to first create this opening, followed by insertion of the bent needle as described above.

   In our experiments, we always introduce full-thickness incisions in the uterine horn. If desired as an alternative methodology for a model of less extensive uterine damage, scraping with the bent 25G needle can be restricted to only the innermost layers of the uterus, such as the endometrium. In addition, we do not suture the uterus closed to avoid introducing foreign-body responses as an additional variable; however, this can be an alternative approach for this procedure.

5. Carefully return the injured uterine horn to the body cavity. See Figure 6A and 6B.

   The surgeon should avoid introducing contact between the open uterine incision and the ovarian fat pad during this process. This can result in adhesions between the fat and the open wound, thus affecting wound healing and introducing an undesirable variable in interpreting the aftereffects of uterine injury.

6. Suture the fascia closed. Remove excess suture thread. See Figure 6C–E.

   A single suture is sufficient to close the incision.

7. Use the AutoClip applicator to close the skin incision. See Figure 6F.

   Two clips may be necessary to close the incision.

8. Inject the animal subcutaneously (S.C.) with 10 mg/kg Carprofen for pain relief. Inject 0.5 mL of saline (0.9% sodium chloride) S.C. as post-operative support to aid in recovery. See Figure 7A.

   As shown in Figure 7A, we suggest a location over the ribs as a site for administering Carprofen and saline since the skin can be stretched taut while scruffing the animal. Carprofen can be injected on one side and saline on the other. Successful subcutaneous injection will appear as a small, liquid-filled distension of the skin. If the underlying fascia has been punctured, no such subcutaneous bubble will form.

9. Place the animal in a heated cage for post-operative recovery (Figure 7B). Use an ear-punch to mark the animal. Monitor the animal until movement resumes. Return the animal to a regular housing cage with bedding once she is able to lift her head. If applicable, perform repeat vaginal lavage of animal to verify estrous staging (see Supporting Protocol 1).

   Full recovery of movement typically occurs within approximately 2–4 hours.

10. Return animal(s) to be housed in the animal facility. Monitor animals daily for signs of pain and provide additional daily injections of Carprofen as needed for pain relief, up to 3 days post-op.

Figure 4:

Incisions to access uterus.

A. Make a small skin incision (~4 mm) dorsally just above the animal’s hind leg.

B. Make a small incision (~4 mm) in the underlying fascia.

C. Grip the underlying fat pad.

D. Pull the fat pad connected to the ovary and uterus out of the body cavity via the incisions generated in (A) and (B).

Figure 5:

Induction of uterine injury.

A. Diagram of needle insertion site for procedure. Diagram created using BioRender.

B. Insert burred needle into top of uterus just below the utero-tubal ligation.

C. Scrape forcefully and repeatedly along the interior of the uterus on the anti-mesometrial surface.

D. Continue scraping until a full-thickness incision is generated along the length of the uterine horn.

Figure 6:

Closure of surgical site.

A. Using forceps, gently return injured uterine horn to body cavity.

B. Return overlying ovarian fat pad to body cavity.

C. Grasp the suture needle using a needle driver.

D. Suture the fascia.

E. Cut off excess suture material.

F. Press skin incision closed and secure with a wound clip.

Figure 7:

Post-surgery care.

A. Inject Carprofen subcutaneously as analgesia. Perform similar injection of saline on the other side of the animal.

B. Place animal in a heated cage until consciousness and movement are regained.

BASIC PROTOCOL 3: Mating and dissection of pregnant mice as endpoint analyses

Introductory paragraph:

Following uterine injury, several different endpoint analyses may be conducted depending on the research question of interest. For example, dissection of non-pregnant animals post-injury can provide macroscopic, cellular, and molecular information about uterine wound healing as a function of estrous phase at time of injury (Zhang et al., 2024). In addition, animals bearing uterine injury may be impregnated either via natural mating or embryo transfer in order to assess the impact of injury on subsequent pregnancy outcomes.

Here we describe the procedure for setting up timed matings and dissection of animals during pregnancy to address embryonic and placental development in a damaged uterine environment. We provide details of dissection at E12.5 of gestation as an example; the user should decide the appropriate gestational age of sacrifice depending on the exact scientific question.

Materials:

Petri dish (e.g. Fisher Scientific, cat. no. 08-757-100)

Forceps (e.g. Fine Science Tools, cat. no. 11255-20)

Scissors (e.g. Fine Science Tools, cat. no. 14060-09)

Probe seeker with bent end (e.g. Fisher Scientific, cat. no. 08-995)

Spray bottle (e.g. Fisher Scientific, cat. no. S413505P)

70% Ethanol (e.g. Fisher Scientific, cat. no. BP82031GAL)

Protocol steps:

1. After post-operative recovery, co-house the injured animal in a cage with a stud male of at least 11 weeks of age.

   Duration of recovery will depend on the researcher’s specific question of interest, such as how length of recovery or active interventions during recovery may impact subsequent pregnancy outcomes. We typically allow for 30 days of recovery (with a range of 5 days to 105 days) prior to co-housing with a male to initiate timed matings.

   If desired, the researcher may seek institutional approval to perform trio or harem matings between a single male and multiple female animal subjects to reduce the cost of housing and increase the efficiency of this experimental approach. Please note that such matings are designed for the intention of dissections during pregnancy; live births resulting from trio or harem matings are not generally permitted due to stress to the animals.

2. Check for the presence of vaginal plugs the following morning. Continue to check each morning until a plug is identified, at which point the female can be housed separately. See Behringer et al. 2016 for further details on checking vaginal plugs (Behringer, Gertsenstein, Nagy, & Nagy, 2016).

   Plugs can appear as either an obvious white mass in the vaginal canal or a bulging of the entire vaginal area. If no sign of a plug is externally visible, gently insert the probe seeker into the vaginal canal. If resistance is encountered prior to full insertion of the probe end into the vagina, a plug is present.

   Plug checks should ideally be performed around 7 am or as early as possible so as to minimize the chance of the plug falling out.

3. Sacrifice the animal at the desired gestational age, with the date of plug corresponding to E0.5.

   Follow your institution’s animal euthanasia guidelines. For example, 8 minutes of CO₂ followed by 2 additional minutes in the euthanasia chamber and cervical dislocation to ensure death, is an appropriate procedure.

4. After confirming euthanasia, place the animal in a supine position. Set up the dissection station with the dissection tools (forceps, scissors) and Petri dishes filled with 1x PBS. Spray the abdomen with either 70% ethanol, water, or 1x PBS to wet the fur and prevent loose strands of fur from adhering to the dissection site. See Figure 8A.

5. While holding the skin of the abdomen with forceps, create an incision in the lower abdomen with the scissors. Continue cutting in a V shape along the length of the abdomen. See Figure 8B–D.

6. While holding the underlying abdominal muscle wall with forceps, create an incision and continue cutting a V shape until the underlying abdominal cavity is exposed. See Figure 8E–F.

7. Move aside the abdominal contents, such as the intestines, until the uterus is exposed. Pull the uterus containing embryos into view. See Figure 8G.

8. Dissect uterus along with enclosed embryos from abdominal cavity. Place uterus into a Petri dish or other container filled with 1x PBS. See Figure 8H.

   Dissection can be performed by first grasping the ovarian fat pad, followed by using scissors to separate the uterus from the rest of the body cavity by severing the ovarian fat, the mesometrial arteries, the vagina and cervix, the mesometrial arteries along the other uterine horn, and lastly, the second ovarian fat pad.

9. Perform further dissection as desired for the appropriate downstream analysis, such as microscopy, histology, or molecular analyses. See Figure 9 for sample data.

Figure 8:

Dissection of pregnant animal.

A. Prepare dissection area with tools and plates containing 1x PBS. Spray euthanized animal’s abdomen with 70% EtOH or other liquid to wet the fur.

B. Grasp the lower abdominal skin with forceps.

C. Cut a V-shaped skin flap in the lower abdomen.

D. Pull back the skin flap to expose the underlying abdominal wall.

E. Cut open the abdominal wall in a V-shape.

F. Pull back the abdominal wall flap to expose the internal organs.

G. Pull out the pregnant uterus for dissection.

H. Place uterus into a 10 cm plate containing 1x PBS for subsequent dissections.

Figure 9:

Examples of expected outcomes following uterine injury.

A. Image of a uterine wound incurred during diestrus, 3 days post-injury. The open wound is indicated with an orange arrow.

B. Image of a uterine wound 1 month post-injury. The prior injury site is indicated with an orange arrow.

C. Image of resorptions within an injured uterine horn.

D. Images of a normal singlet placenta and embryo.

E. Image of a close pair of misspaced embryos with their placentas fused together.

F. Image of a close pair of misspaced embryos in which one embryo is underdeveloped (orange arrow) due to embryo crowding.

SUPPORT PROTOCOL 1: Estrous staging of animals

Introductory paragraph:

Certain downstream effects of uterine injury are dependent upon estrous cycle stage at the time of surgery; for instance, embryo spacing defects occur in pregnancies that follow diestrus but not estrus injuries, while wound healing dynamics differ between diestrus vs. estrus injuries (Zhang et al., 2024). Furthermore, given the dramatic effects of estrous on both the structure as well as cellular and molecular composition of the uterus, it is important that the same estrous phase is chosen for reliable comparisons across samples. For example, in dissecting non-pregnant animals to assess wound healing or in preparation for molecular approaches such as RNA-seq, the same estrous phase should be chosen for comparison (Bertolin & Murphy, 2014; Roberson et al., 2022; Zhang et al., 2024). Therefore, determination of estrous phase can be a critical step for any experimental manipulation and downstream molecular or histological analysis. The most reliable method for assessing estrous cycle stage in live mice is vaginal smear cytology (McLean, Valenzuela, Fai, & Bennett, 2012), which can be performed using the steps described below.

Materials:

Glass staining dish for slides (e.g. Fisher Scientific, cat. no. 08-817)

Superfrost Plus microscope slides (Fisher Scientific, cat. no. 22-037-246)

50 × 24 mm Cover Slips (Fisher Scientific, cat. no. 12-541-055)

Sterile distilled water (e.g. Gibco, cat. no. 15230170)

0.1% crystal violet solution (see Reagents and Solutions)

200 μL micropipette

Transfer pipette (e.g. Thermo Scientific Samco Standard Disposable Transfer Pipettes, cat. no. 13-711-9D)

Protocol steps:

1. Perform vaginal lavage with 50–100 μL of sterile H2O, pipetting in and out approximately 5 times using a 200 μL micropipette.

2. Spread lavage contents onto glass slide.

3. Allow to dry.

   This should take approximately 30 min. This can be accelerated by placing slides on a slide warmer at 37 deg C.

4. Stain with 0.1% crystal violet solution. Placed into a glass Coplin jar with the solution for ~1 minute.

   Alternatively, to conserve crystal violet solution, place slides flat on multiple layers of paper towels. Then, using a disposable transfer pipette (e.g. Thermo Scientific Samco Standard Disposable Transfer Pipettes, cat. no. 13-711-9D) or a 1000 uL pipette, transfer ~0.5 mL of crystal violet solution to cover the surface of the slide. Allow to sit for ~1 minute, then remove solution by pouring off onto paper towels. Immediately proceed to next step.

5. Place stained slides into a glass Coplin jar with distilled water for ~1 minute to wash.

6. Remove slides from Coplin jar and mount by placing a coverslip over the surface of the slide. Dab off excess liquid from the edges with a paper towel.

7. View under a microscope to assess estrous stage. See Figure 10 for examples of vaginal cytology patterns and their corresponding phase of estrous.

Figure 10:

Determination of estrous cycle stage by vaginal cytology.

Cytology of vaginal lavage smears stained with crystal violet showing cell types indicative of the proestrus, estrus, metestrus, and diestrus phases of the mouse estrous cycle.

We strongly recommend taking a second vaginal cytology sample at the completion of the procedure to confirm that the animal has remained in the same phase. Mice can occasionally cycle quickly and enter a different phase at the time of the procedure.

Additional information regarding the assessment of estrous phase by vaginal cytology can be found in McLean et al. 2012 (McLean et al., 2012).

REAGENTS AND SOLUTIONS:

Recipes for reagents or solutions that are unique to the protocols described in the manuscript. Provide details on how to produce these recipes and include storage information (temperature and duration).

Preparation of 0.1% crystal violet solution

1. Dissolve 1 g of crystal violet (Sigma-Aldrich, cat. no. C0775) in 50 mL of 100% ethanol (e.g. Fisher Scientific, cat. no. BP2818100).

2. Adjust total volume to 1 L with distilled or ultrapure water (e.g. Gibco, cat. no. 15230170).

3. Store at room temperature.

Preparation of ketamine/xylazine solution (10 mg/mL ketamine + 1 mg/mL xylazine in 0.9% saline)

1. Remove and discard 1.1 mL of 0.9% sodium chloride from a 10 mL vial.

2. Inject 0.1 mL of 100 mg/mL Xylazine HCl and 1 mL of ketamine HCl into 0.9% vial.

3. Label bottle with bottle number, contents, and date.

4. Store at room temperature in a locked drawer for up to one month in compliance with institutional regulations for controlled substances.

COMMENTARY

BACKGROUND INFORMATION:

Although Cesarean sections (C-section) account for ~33% of deliveries in the US and up 55% of deliveries in countries such as China and Brazil, little is known about how the uterus heals after injury and how these injuries impact subsequent pregnancy outcomes. While clinical and epidemiological evidence implicates prior C-section as a major risk factor for pregnancy disorders such as placenta accreta spectrum (PAS) disorder and placenta previa, the mechanisms by which these disorders arise are unclear. As a result, diagnostics and treatments remain inadequate, contributing to substantial maternal and fetal morbidity and even mortality. With increasing rates of C-section in the US, the prevalence of PAS continues to rise.

In spite of these trends, animal models for understanding the structural, cellular, and molecular consequences of uterine injury remain limited. Recently, Buhimschi et al. examined uterine wound healing following C-section in mice (Buhimschi et al., 2010). In addition, Burke et al. and Li et al. demonstrated reproductive defects in mice that experienced prior uterine injury (Burke et al., 2020; Li et al., 2020). As with our model, Burke et al. and Li et al. both performed full-thickness uterine incisions and assessed subsequent pregnancy outcomes. Overall, while many aspects of the methodologies and resulting findings described in these 3 studies and in ours (Zhang et al., 2024) are similar, we also outline some key differences below.

The methodology presented in this protocol provide several advantages. First, by accessing the uterus dorsally as opposed to abdominally via C-section as described by Buhimschi et al. and Burke et al., our procedure allows for smaller surgical incisions, faster procedure time, and rapid recovery of the animal, potentially enabling broader adoption of our approach. In addition, in contrast to Li et al., since no suturing of the uterus is required in our approach, the wounds heal independently of suturing technique, thus allowing for greater consistency between animals while also enabling investigation of spontaneous uterine wound healing without confounding foreign body responses. In addition to the use of suturing, Li et al. used a slightly different methodology to introduce uterine injury: a cut using ophthalmic scissors as opposed to the burred needle employed in our approach.

As a result of either these different methodologies and/or the strains of mice used, some but not all of the outcomes we identified in our model were also observed in these other studies. For example, both we and Li et al., but not Burke et al., noted embryo spacing and embryo resorption. Burke et al., Li et al., and we (Zhang et al.) identified invasive placentas. Overall, we present here a novel and facile methodology for inducing mechanical uterine injury in mice to investigate uterine wound healing and in utero development with parallels to relevant human pregnancy disorders.

CRITICAL PARAMETERS AND TROUBLESHOOTING:

Several factors should be considered prior to conducting these protocols. First, it is important to calculate the number of animals required for these experiments and to plan ahead with ordering or breeding to obtain the desired numbers. It is especially critical to ensure sufficient numbers of the desired age range, as well as to ensure sufficient numbers of animals in the desired stage(s) of estrous. Female mice of 7–8 weeks of age are recommended for studies pertaining to reproductive defects, whereas pre-pubertal or aged mice may be used for other experimental questions. Secondly, as mentioned in Basic Protocol 1, the strain of mice used can impact both the surgical procedure itself as well as the experimental outcomes. For instance, C3H mice typically require higher doses of ketamine/xylazine than C57BL/6 or BALB/c mice. Adjustments should be made accordingly in order to achieve a surgical plane of anesthesia. An appropriate plane of anesthesia with ketamine/xylazine is achieved when the forelimb reflex is entirely absent and the hindlimb reflex to light touch is no longer present.

Certain problems can arise during the course of these experiments. Here we present two common problems, their causes, and potential solutions in the Table 1.

Table 1.

Troubleshooting Guide.

Problem	Cause	Solution(s)
Severing the uterus at the uterotubal ligation	Excessive force applied to forceps used to secure the uterus And/or excessive injury to the uterus during needle insertion	First make a small hole with a straight, unmodified 25G, 27G, or 30G needle, followed by insertion of the burred 25G injury tool to incur damage
Adhesions forming between ovarian fat pad and open uterine incision, which impacts interpretation of wound healing results	Contact between ovarian fat pad and uterine incision	Exercise care when returning the injured uterine horn into the body cavity to avoid contact with ovarian fat pad
Surgical complications	Wound not fully closed, infection, bleeding	Use a wound clip remover to remove and reapply any surgical clips that do not fully close the wound; check animals daily to check for signs of pain or distress and administer supportive care, or if necessary, euthanasia, in a timely manner according to the researcher’s animal ethics committee recommendations

UNDERSTANDING RESULTS:

Immediately following surgery, animals typically are partially mobile within ~1 hr. and fully mobile within ~4–6 hours. Welfare checks are performed on these animals daily to monitor for signs of pain and until the skin incision has healed and the skin clips are ready to be removed. Complications from surgery are typically noted shortly after surgery, such as skin incision dehiscence or pain.

After surgery, animals can be sacrificed at various timepoints to assess uterine wound healing. Examples of uterine wound healing are shown in Figure 9, illustrating damage to a diestrus-injured uterus 3 days post-injury (Fig. 9A) and minimal scarring to uteri after >1 mo. of recovery (Fig. 9B). Further interventions, such as drug treatments during surgery or recovery, or additional surgeries, are also possible. Animals can be subjected to matings or embryo transfers to examine the impact of uterine injury on embryo implantation and development. Injury often leads to effects such as uterine bleeding and embryo resorptions (Fig. 9C) and embryo misspacing (Fig. 9E vs. Fig. 9D), which often leads to crowding for resources and resorption of one of the two misspaced embryo pairs.

For a comprehensive description of anticipated results in uterine wound healing and subsequent pregnancy outcomes, please refer to Zhang et al. 2024 Biol Reprod (Zhang et al., 2024).

TIME CONSIDERATIONS:

Estrous cycle staging requires approximately 1–2 hours of time to perform vaginal lavage collection, drying of microscope slide, crystal violet staining, and imaging.

Uterine injury and sham surgeries require approximately 30–45 minutes per animal, including approximately 15 minutes to achieve a surgical plane of anesthesia, and 15–30 minutes to perform the surgery itself.

Immediate recovery from surgery requires at least 1 hr. before animals are partially mobile and able to lift their heads sufficiently for return to their cages.

Recovery from the incurred injury is dependent upon the specific research question. For our work as described in Zhang et al. 2024 Biol Reprod (Zhang et al., 2024), recovery periods ranged from 5 days to >4 months.

SIGNIFICANCE STATEMENT:

Approximately 1 in 3 deliveries in the US are via Cesarean section (C-section), and yet how the uterus heals and how an injured uterus leads to various pregnancy complications is unclear. Complications arising from a history of uterine injury carry substantial disease burden and include uterine rupture and scarring, as well as pregnancy disorders like placenta previa and placenta accreta. An animal model is thus critical for unraveling the mechanisms of uterine wound healing and its impacts on pregnancy outcomes, as well as for enabling the development of diagnostics and therapeutics. Here we describe a procedure for incurring uterine injury in a mouse model, which presents an important step forward in understanding uterine damage and its associated pregnancy disorders.