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Method Article

Mouse Model of Surgical Uterine Injury and Subsequent Pregnancy Outcomes

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DOI:

10.3791/67977

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June 27th, 2025

 , 
1Department of Physiology and Membrane Biology, University of California, Davis, 2Department of Obstetrics and Gynecology, University of California, Davis

In This Article

Summary

This protocol describes a procedure for inducing uterine injury in a mouse model, followed by downstream analyses during a subsequent pregnancy.

Abstract

Uterine injury from Cesarean sections (C-sections) and other procedures (dilation and curettage, myomectomies, etc.) can lead to several morbidities in subsequent pregnancies, including placenta previa, placenta accreta spectrum (PAS), Cesarean scar pregnancy, and uterine rupture. C-sections account for approximately 30% of deliveries in the US, with rates projected to rise. Therefore, a deeper understanding of the mechanisms by which injury-associated pregnancy disorders arise and a platform for testing possible interventions are essential. To achieve these objectives, there is a critical need for animal models of uterine injury, particularly those that address the impact of injury on subsequent in utero outcomes. This protocol describes a novel surgical procedure for a model of mechanically induced uterine injury in the laboratory mouse (Mus musculus). Steps taken in preparation for surgery, induction of the uterine injury itself, and post-surgical recovery procedures are demonstrated. Additional information is provided regarding the downstream dissection of both non-pregnant and pregnant mice. Overall, by detailing a set of procedures for inducing uterine injury in an in vivo model, this protocol presents a tractable method for elucidating the molecular and cellular events of uterine wound healing and its effects on subsequent pregnancy.

Introduction

Although 1 in 3 deliveries in the US are conducted via Cesarean section (C-section), little is known about how the uterus heals and how an injured uterus leads to complications in subsequent pregnancies1,2,3,4,5,6,7,8,9,10,11,12,13. These complications include uterine rupture, placenta previa, PAS, and Cesarean scar pregnancy, which together affect ~2.5% of patients who have had one or more C-sections1,2,6,8,12,14,15,16,17. Placenta previa is characterized by the abnormal placement of the placenta in the lower portion of the uterus4,16. This not only causes physical obstruction of the cervix and an elevated risk of bleeding during pregnancy, but is also a significant risk factor for the more severe, invasive condition of PAS8,12. PAS is defined by over-invasion of the placenta too deeply into maternal tissues, even as far as the bladder or rectum, and often requires a C-section hysterectomy to preclude life-threatening maternal hemorrhage12,17. Cesarean scar pregnancies, though rare, carry a high rate of related morbidities, including miscarriage, fetal death, and preterm birth18,19. In addition to complications associated with a history of prior C-sections, other uterine procedures such as myomectomy or dilation and curettage (D&C) also lead to patient morbidities. For instance, ~30% of patients undergoing a D&C after a late spontaneous abortion and up to 46% of patients undergoing a second hysteroscopic resection for leiomyoma develop intrauterine adhesions characteristic of Asherman syndrome (AS)20,21. AS impacts ~2% of females, causing amenorrhea, dysmenorrhea, embryo implantation failure, recurrent pregnancy loss, preterm labor, and/or PAS21,22. Therefore, given the substantial major morbidities and mortalities associated with prior uterine trauma, an experimental animal model is critical not only for elucidating the mechanisms by which uterine wounds heal and how uterine injury impacts subsequent pregnancy outcomes, but also as a platform for the development of novel diagnostics and therapeutics23,24.

Several studies have thus developed mouse models of uterine injury. However, early studies either did not examine the consequences of uterine injury on subsequent pregnancies25 or limited the damage to the uterine luminal surface to investigate embryo implantation outside the window of endometrial receptivity in ovariectomized mice26. To address these knowledge gaps, the model described in this protocol, along with other recent studies, presents a novel surgical methodology for introducing uterine injury into mice27,28. This protocol uses the common laboratory mouse as the model organism due to its wide accessibility, genetic tractability, short gestation, rapid time from birth to puberty, large litter sizes, and cost-effectiveness. In addition, the anatomical feature of two uterine horns in mice allows for an internal control with an uninjured horn in each experimental animal, as well as the ability to separate local defects directly at the scar vs. global defects throughout the entire uterine environment. The approach described in this Protocol entails a simple dorsal laparotomy, followed by the use of a burred needle to create an incision through both the endometrial and myometrial layers of a single uterine horn. This methodology most closely mimics injuries seen during C-sections and myomectomies, though it can easily be modified to investigate non-incisional injuries such as D&Cs, which precede AS29,30. Following injury, animals are allowed to recover and then mated with stud males for timed pregnancies to examine the consequences of injury on subsequent in utero outcomes27,28.

Although similar in overall strategy to other recent studies, there are aspects of this protocol that provide distinct advantages. For instance, both Li et al. and Burke et al. accessed the uterus abdominally as opposed to dorsally as described here31,32. A dorsal approach is both simpler to perform and more conducive to rapid post-operative recovery of the animal. For the non-pregnant mice in the Burke et al. study, bent needles or curettes were used to scrape the endometrium, thus more closely mimicking D&Cs, as opposed to the full-thickness incisional injuries in this Protocol that more closely model C-sections and myomectomies32. In peripartum mice, Burke et al. introduced incisions on the mesometrial as opposed to the anti-mesometrial surface where embryos implant, as described here. By using a burred needle as opposed to scissors used by Li et al.31, this approach can more easily be modified from full-thickness to endometrium-restricted injuries to model both D&Cs as well as C-sections and myomectomies. Both Li et al. and Burke et al. closed uterine wounds with sutures, which, though more faithful to the events during a C-section, introduces additional variability into the healing process as well as the potential for foreign body effects31,32,33,34,35,36. Similar studies in rats have used abdominal routes to generate full-thickness incisional37,38 or excisional39 uterine wounds that are closed using sutures37,38,39 in both non-pregnant38,39 and pregnant37 animals. Therefore, the protocol presented here is particularly suitable for researchers interested in a facile and reproducible approach that requires minimal surgical experience, addresses embryo implantation- and placentation-related events following a full-thickness uterine injury, and is easily adaptable to a wide variety of questions, including D&C-like injuries to the endometrium27,28.

Step 1 of this protocol outlines pre-operative steps. Step 2 describes the surgical procedure for inducing injury to a uterine horn. Finally, Step 3 details the dissection of a pregnant dam to investigate the effects of uterine surgery on a subsequent pregnancy. Altogether, this study presents a unique rodent surgical model of uterine wound healing and associated pregnancy outcomes that is quick to perform and easy to adopt.

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Protocol

All procedures demonstrated in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) of the University of California, Davis (IACUC Protocol #23985). The procedures described in this protocol are typically performed on C3H females between 7-10 weeks of age and 19-25 g in weight, though this protocol is suitable for female mice of all strains, ages, and weights, depending on the researcher's question. Monitor animals prior to surgery to ensure that only healthy animals with no signs of injury or distress are used for studies. The details of the reagents and the equipment used are listed in the Table of Materials.

1. Preparation of the surgical area

  1. Prior to the day of surgery, place all surgical instruments (fine forceps (#5 or #55), needle holder, curved iris scissors, and skin clip applicator (loaded with clips)) in sterilization pouches and autoclave.
  2. If using an injectable anesthetic, prepare a working dilution of 10 mg/mL ketamine and 1 mg/mL xylazine in 0.9% sodium chloride saline.
    NOTE: Alternatively, isoflurane delivered via nosecone is an acceptable form of anesthesia40.
  3. Immediately prior to surgery, don personal protective equipment (PPE): lab coat, surgical hair cap, surgical facemask, and gloves.
  4. Prepare the pre-operative area.
    NOTE: This may consist of one absorbent bench pad lined with paper towels and a cage in which the animal may rest after receiving anesthesia prior to surgery. Line the animal cage with paper towels and place it next to the pre-op bench pad.
  5. Weigh the animal designated for surgery in a tared plastic beaker or similar container atop a scale (Figure 1A).Record the weight to calculate the appropriate amounts of anesthetic, analgesic, and saline.
  6. Administer injectable anesthesia to the animal via the following steps:
    1. Fill an insulin syringe with the appropriate volume of the working dilution of ketamine/xylazine (see step 1.1) to achieve a dose of 100 mg/kg ketamine and 10 mg/kg xylazine (Figure 1B).
      NOTE: For example, an animal weighing 22 g would require 0.22 mL of the 10 mg/mL ketamine and 1 mg/mL xylazine anesthetic solution.
    2. Inject the appropriate dose of ketamine/xylazine intraperitoneally (I.P.) after securely scuffing and inverting the animal into a supine position41 (Figure 1C).
      NOTE: A scuff guard may be used as needed.
  7. Monitor the animal for approximately 3 min. Inject an additional 0.02 mL of ketamine/xylazine solution I.P.41 if movement is observed during this window of time.
  8. Check for adequate sedation by performing a front toe pinch ~10 min after anesthesia is delivered40,42.
    NOTE: Animals typically achieve a surgical plane of anesthesia within 10-15 min. C3H mice typically require ~20% more anesthetic than C57BL/6 or BALB/c animals (e.g., 0.24 mL ketamine/xylazine solution for a 20 g animal). Dosing may need to be determined empirically based on each individual strain.
  9. After cessation of movement, subcutaneously (S.C.) inject41 the animal with 0.5 mg/kg extended-release Buprenorphine (Figure 1D).
    NOTE: Given the small volume and viscous nature of the extended-release Buprenorphine, injecting it near the shoulder blade while tenting the skin with the other hand is recommended.
  10. To protect the animal's eyes, apply ophthalmic ointment using a sterile cotton swab (Figure 2A).
  11. Place the anesthetized animal onto the pre-operative absorbent pad lined with paper towels.
  12. Carefully shave the incision site at the lower to mid back using a fur trimmer (Figure 2B). Remove excess fur with the use of paper towels, followed by a lint roller (Figure 2C).1.12
    NOTE: Hair removal cream may be applied using a sterile cotton swab for complete fur removal.
  13. Disinfect the incision site using alcohol prep pads and povidone-iodine swabsticks in the following manner:
    1. Starting at the center of the intended incision site, wipe outward with the alcohol prep pad in a circular motion (Figure 2D).
    2. Next, wipe the incision site with a povidone-iodine swabstick in a similar fashion (Figure 2E).
    3. Alternate between fresh alcohol prep pads and povidone-iodine swabsticks for a total of 3 rounds.
  14. Prepare the surgical area as follows (see Figure 3A):
    1. Place a second absorbent pad under an illuminator, followed by a sterile surgical drape over the absorbent pad.
    2. Sanitize the illuminator with 70% ethanol. Change to sterile gloves before handling the sterilized instruments and making the incision.
    3. Place sterile surgical tools at the edges of the surgical drape and place the animal in the center of the surgical area.
  15. Create a sterile field by placing a sterile drape (e.g. Glad Press 'n Seal) over the animal, leaving only the snout exposed (Figure 3B).
    NOTE: During use, clean surgical tools may be placed on sterile surfaces such as the sterile drape within the surgical area.
  16. Cut a small square over the shaved region in preparation for generating an incision at that site (Figure 3C,D). See NOTE for step 2.1 (below) for the specific location of the incision site.

2. Surgical induction of uterine injury

NOTE: This section describes the surgery and procedure for introducing injury into a single uterine horn of a non-pregnant female mouse. First, a dorsal laparotomy is performed to expose the top of the uterine horn. Then, a burred 25 G needle is inserted into the uterine lumen to damage the uterine endometrium and, if desired, to cut through the myometrium. This creates a longitudinal wound in the injured uterine horn, enabling further experimentation to address questions such as how the uterus heals and how injury impacts a subsequent pregnancy. It is important that the animal is properly sedated before performing the surgery. As described in Step 1, several parameters indicate that an animal is adequately anesthetized. These include the cessation of movement, a reduction in respiratory rate, and the absence of a forelimb reflex. In addition, the hindlimb reflex upon a light touch should be absent, though this reflex sometimes remains present despite adequate anesthesia if sufficient pressure is applied40,42.

  1. Cut a 2-3 mm incision in the mouse dorsal skin using a pair of scissors (straight or curved iris scissors) while holding the skin with fine forceps (#5 or #55).
    NOTE: Cut ~⅓ distance between the base of the tail and neck and ~0.5 cm from the midline. See Figure 4A.
  2. Cut a 2-3 mm incision in the fascia immediately overlaying the ovarian fat pad (Figure 4B).
  3. Gently grip the ovarian fat pad with a pair of forceps and extract it from the incisions in the fascia and skin (Figure 4C).
    NOTE: Parts of the fat pad may need to be cut so that the ovary and uterus can be removed from the body cavity. Avoid injuring any part of the female reproductive tract itself, as well as the spleen and any large blood vessels near the fat pad.
  4. After exposing the upper third of the uterus, perform the following (Figure 4D):
    1. Gently secure the uterotubal ligation with forceps (Figure 5A,B).
    2. Use the sharp end of a burred needle to create a small hole in the top of the uterus (Figure 5A,B).
    3. Insert the needle carefully and begin scraping the anti-mesometrial surface of the uterine horn (Figure 5C).
    4. Continue scraping until complete perforation of both the endometrial and myometrial layers has been achieved (Figure 5D).
      NOTE: Excessive pressure with forceps and/or needle insertion can rip the uterus at the uterotubal ligation, which results in the separation of the uterine horn from the oviduct and ovary. If it is difficult to create an entry hole into the uterus with the burred needle without damage to the uterotubal ligation, an unburred 25 G or 27 G needle may be used as an alternative. The burred needle can then be inserted into this hole to generate the incision as described above. In the approach described here, full-thickness incisions are introduced in the uterine horn. However, if less extensive uterine damage is desired, the burred 25 G needle may be used to scrape only the innermost layers of the uterus, such as the endometrium. In addition, this procedure does not use sutures to close the uterus to avoid introducing foreign-body responses as an additional variable; however, uterine suturing is an acceptable alternative for this procedure.
  5. Gently return the injured uterine horn to the body cavity using forceps (Figure 6A,B).
  6. Suture43 the fascia closed with size 4-0 suture and remove any excess suture (Figure 6C-E).
    NOTE: A single suture is typically sufficient to close the incision in the fascia.
  7. Close the skin incision with clips using the skin clip applicator (Figure 6F).
    NOTE: One to two clips are necessary to close the incision.
  8. Subcutaneously inject41 10 mg/kg Carprofen as analgesia and 0.5 mL of saline (0.9% NaCl) for post-operative supportive care. See Figure 7A.
  9. Use an ear-punch or numbered metal ear tags to identify the animal44.
  10. Place the animal in a heated cage for post-operative recovery (Figure 7B).
    NOTE: Once the animal is able to lift her head, return her to a regular housing cage with bedding. See step 2.12 below.
  11. If performing additional surgeries with the same surgical tools, then perform the following steps:
    1. First, clean with 70% ethanol and paper towels.
    2. Then, resterilize the tools by placing them in a dry glass bead sterilizer for 30 s.
    3. Allow the tools to cool to room temperature before further use.
  12. Return animal(s) to be housed in the animal facility after resumption of movement, ~1-2 hrs following recovery from surgery if ketamine/xylazine were used or ~5 min following anesthesia with isoflurane.
  13. Monitor animal(s) daily for signs of pain and distress and provide daily injections of 10 mg/kg carprofen as needed for up to 3 days post-surgery.
    NOTE: The grimace scale45 and body condition scoring46 are standard veterinary assessments of animal well-being.

3. Mating and dissection of pregnant mice to assess consequences of uterine injury

NOTE: Depending on the specific research question, various endpoint analyses may be performed post-uterine injury. For example, non-pregnant animals may be dissected at different timepoints post-injury to examine the kinetics of wound healing27. In addition, injured animals may be impregnated via natural mating or embryo transfer to examine the impact of injury on subsequent pregnancies27. This Step describes the assessment of embryonic and placental development in a damaged uterine environment. In particular, this Step provides details on setting up timed matings and dissection of animals at embryonic day 12.5 of gestation (E12.5, with the morning of vaginal plug defined as E0.5; more information on identifying vaginal plugs in step 3.2 below). Altogether, Step 3 provides an example of downstream analyses that can be conducted post-injury to examine a specific set of experimental questions.

  1. After post-operative recovery, perform timed matings47 by co-housing the injured female with a stud male who is 11 weeks of age or older.
    NOTE: The length of recovery post-injury will be determined by the specific research question. Studies using this Protocol typically allow for 30 days of recovery (with a range of 5-105 days tested) prior to initiating timed matings. Trio or harem matings reduce the cost of housing and increase the efficiency of this experimental approach. However, please note that such mating configurations require institutional approval and are intended solely for the purposes of dissections during pregnancy; live births from trio or harem matings cause stress to the animals and are not generally advised or permitted.
  2. Perform vaginal plug checks each morning until a plug is identified (see Figure 8), at which point the female can be housed separately from the stud male.
    NOTE: Plugs will appear as either a white mass in the vaginal canal (Figure 8A) or a bulging of the entire vaginal area. If no sign of a plug is externally visible, a probe seeker may be inserted slowly into the vaginal canal (Figure 8C,D). Any resistance before full insertion of the tip up to the bend in the probe seeker is an indication of a plug. Plug checks should be performed at ~7 am or as early as possible in the morning since plugs fall out over the course of the day. For more details on checking vaginal plugs, see Behringer et al.48. For additional and more reliable confirmation of pregnancy, high-frequency ultrasound can be used49,50. If ultrasound is unavailable or undesired, weight gain has been demonstrated to predict pregnancy with reasonable accuracy by E7.7551.
  3. Sacrifice the animal (following institutionally approved protocols) at the desired gestational age. See the NOTE below for details on how to perform animal euthanasia and how to select a gestational age for dissection..
    NOTE: The plug date is designated as E0.5. Choose a gestational age at the time of sacrifice based on the experimental question being addressed. For example, E4.5-E5.5 is appropriate for investigating peri-implantation biology. E11.5-E14.5 is advisable for investigating mid-gestational placental defects, including the presence of fused placentas. Follow institutional guidelines for animal euthanasia. An acceptable procedure is an 8-min CO2 incubation period, followed by a 2-min waiting and observation period, and finally, secondary confirmation of euthanasia via cervical dislocation.
  4. After euthanasia, prepare for dissection as follows (see Figure 9A):
    1. Place the animal in a supine position on a dissection surface.
    2. Obtain the necessary dissecting tools (forceps, scissors), along with Petri dishes filled with 1× PBS.
    3. To prevent loose strands of fur from adhering to the dissection site, first wet the abdominal fur with either 70% ethanol, water, or 1× PBS prior to dissection.
  5. Make an incision in the lower abdomen by holding the skin of the abdomen with forceps and cutting a V shape along the length of the abdomen with scissors until reaching the bottom of the ribcage (~2 cm incision for each stroke of the letter "V"). See Figure 9B-D.
  6. Create a V-shaped incision in the underlying abdominal muscle wall (~2 cm incision for each stroke of the letter "V") while holding it with forceps and cutting in a V shape until the abdominal cavity is visible. See Figure 9E,F.
  7. Expose the uterus by gently moving aside the intestines and other abdominal contents. Pull the uterus containing embryos out of the abdominal cavity and into view. See Figure 9G.
  8. Remove the uterus along with the enclosed embryos from the abdominal cavity. Place the uterus into a Petri dish prefilled with 1× PBS. See Figure 9H.
    NOTE: Use forceps to first grasp the ovarian fat pad. Then, use scissors to separate the uterus from the rest of the body cavity by first severing the ovarian fat, followed by the mesometrial arteries, the vagina, the mesometrial arteries along the other uterine horn, and lastly, the second ovarian fat pad. Further dissection may be conducted as needed for the desired downstream analysis, such as microscopy or histology. See Figure 10 for example data.

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Results

Following surgery with the use of ketamine/xylazine as the anesthetic, animals are typically partially mobile within ~1 h. and fully mobile within ~4-6 h. Recovery from isoflurane anesthesia typically occurs within minutes. Any complications that may arise, such as wound dehiscence or pain, are typically evident within 1-3 days following surgery. Perform daily welfare checks to monitor for pain or complications until the skin incision has healed and the skin clips are ready to be removed.

Anim...

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Discussion

C-sections account for ~33% of deliveries in the US, and up to 55%-65% of deliveries in countries such as Brazil and Egypt12,52, and yet little is known about how the uterus heals following injury or how these injuries impact subsequent pregnancies. While prior C-section is a major risk factor for pregnancy disorders like placenta accreta spectrum (PAS) and placenta previa, the mechanisms by which these arise are unclear3,

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Disclosures

The authors have no conflicts of interest to declare.

Acknowledgements

We thank the UC Davis Teaching and Research Animal Care Services (TRACS) for their ongoing husbandry, procedural, and logistical support. We thank the UC Davis Cardiovascular Research Institute (CVRI) Animal Model Core and core director Dr. Yi-Je (Jay) Chen, D.V.M., Ph.D. for technical guidance and support as well as surgical suite use for filming. We thank all members of the Zhang lab for their support of this work. Figure 5A was created with BioRender.com (https://BioRender.com/ig9sfon).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
0.9% NaCl for injection, sterileFresenius Kabi63323-186-10
25 G x 5/8 in. BD PrecisionGlide™ NeedleBD305122
70% ethanolFisher ScientificBP82031GAL
Absorbable 4-0 vicryl sutureEthiconJ214
Animal cagesVWR10712-110 and 10715-323Cages provided by the researcher's animal facility are acceptable
Animal ear-punchFisher Scientific13-812-201
AutoClip wound closing systemFine Science Tools12020-00
U-100 insulin syringeFisher Scientific14-826-79BD brand
Betadine swabsticksMedlineMDS093901
Buprenorphine Extended Release, 1.3 mg/ml (Ethiqa XR)Fidelis Pharmaceuticals099114Alternative extended release Buprenorphine or Buprenorphine HCl are acceptable
Carprofen (RIMADYL) 50 mg/ml ZoetisRIM-00284R1
Curved iris scissorsWorld Precision Instruments501759G
Dry glass bead surgical tool sterilizerVWR INTERNATIONAL75999-324
Dry glass bead refillVWR INTERNATIONAL75999-332
Female mice, typically 8 to 12 weeks of age, e.g., strain C3HCharles River025
Fine scissors - curved 10.5cmFine Science Tools14370-23
Fur trimmerFisher Scientific50-195-4544
Glad Press ‘n SealFisher ScientificNC1089054
Gloves, nitrileFisher Scientific19-130-1597C
H2O, sterile distilledFine Science Tools15230170
Illuminator for surgeriesAmScopeHL150-AY
Ketamine, 100 mg/mL (Zetamine)Dechra501072
Lab coatUlineS-15376W-50
Light microscopeFisher ScientificS13039
Lint rollerFisher Scientific17-018-329
Nair Hair Removal CreamTarget049-06-3304
Needle holderFine Science Tools12500-12
Paper towelsScott1804
Petri dishFisher Scientific08-757-100
Phosphate-buffered saline (PBS)Thermo Fisher Scientific10010023
Plastic beakerThermo Fisher Scientific12010250
Probe seeker with bent endFisher Scientific08-995
Fisherbrand™ Instant Sealing Sterilization Pouches 9 cm (W) x 13.3 cm (H)Fisher Scientific181250
Fisherbrand™ Instant Sealing Sterilization Pouches 9 cm (W) x 23 cm (H)Fisher Scientific181251
Forceps, Fine, #5Fine Science Tools11254-20
Forceps, Fine, #55 Fine Science Tools11295-51
ScaleFisher ScientificS93805
Scissors, surgicalFine Science Tools14060-09
Spray bottleFisher ScientificS413505P
Sterile alcohol prep padsFisher Scientific22-363-750
Sterile ophthalmic veterinary ointmentPuralubePH-PURALUBE-VET
Sterile swabs for eye ointment applicationFisher Scientific18-366-472
Sterilization pouchesFisher Scientific19-910-673
Surgical drapeFisher Scientific50-209-1792
Surgical hair capUlineS-10480BLU
Surgical masksFisher Scientific18-048-010
Thomas Scientific Absorbent Bench Underpad, Bench pads, 16.5 x 23 in./42 x 58cmFisher ScientificNC1588452
Warming/induction chamber with full floor warmingFisher Scientific14-370-302
Wound clip refillsFine Science Tools12022-09
100mg/ml xylazineDechraRompun 100 mg/mL Xylazine

References

  1. Bowman, Z. S., et al. Risk factors for placenta accreta: A large prospective cohort. Am J Perinatol. 31 (9), 799-804 (2014).
  2. Miller, D. A., Chollet, J. A., Goodwin, T. M. Clinical risk factors for placenta previa-placenta accreta. Am J Obstet Gynecol. 177 (1), 210-214 (1997).
  3. Solheim, K. N., et al. The effect of cesarean delivery rates on the future incidence of placenta previa, placenta accreta, and maternal mortality. J Matern Fetal Neonatal Med. 24 (11), 1341-1346 (2011).
  4. Biswas, R., Sawhney, H., Dass, R., Saran, R. K., Vasishta, K. Histopathological study of placental bed biopsy in placenta previa. Acta Obstet Gynecol Scand. 78 (3), 173-179 (1999).
  5. Cramer, S. F., Heller, D. s Placenta accreta and placenta increta: An approach to pathogenesis based on the trophoblastic differentiation pathway. Pediatr Dev Pathol. 19 (4), 320-333 (2016).
  6. Khong, T. Y., Robertson, W. B. Placenta creta and placenta praevia creta. Placenta. 8 (4), 399-409 (1987).
  7. Khong, T. Y., Cramer, S. F., Heller, D. S. Chorion laeve accreta - Another manifestation of morbid adherence. Placenta. 74, 32-35 (2018).
  8. Silver, R. M., Barbour, K. D. Placenta accreta spectrum: Accreta, increta, and percreta. Obstet Gynecol Clin North Am. 42 (2), 381-402 (2015).
  9. Stanek, J., Drummond, Z. Occult placenta accreta: The missing link in the diagnosis of abnormal placentation. Pediatr Dev Pathol. 10 (4), 266-273 (2007).
  10. Tantbirojn, P., Crum, C. P., Parast, M. M. Pathophysiology of placenta creta: The role of decidua and extravillous trophoblast. Placenta. 29 (7), 639-645 (2008).
  11. Zhang, E. T., et al. PRG2 and AQPEP are misexpressed in fetal membranes in placenta previa and percretadagger. Biol Reprod. 105 (1), 244-257 (2021).
  12. Wortman, A. C., Alexander, J. M. Placenta accreta, increta, and percreta. Obstet Gynecol Clin North Am. 40 (1), 137-154 (2013).
  13. Wu, S., Kocherginsky, M., Hibbard, J. U. Abnormal placentation: Twenty-year analysis. Am J Obstet Gynecol. 192 (5), 1458-1461 (2005).
  14. Chazotte, C., Cohen, W. R. Catastrophic complications of previous cesarean section. Am J Obstet Gynecol. 163 (3), 738-742 (1990).
  15. Jurkovic, D., et al. First-trimester diagnosis and management of pregnancies implanted into the lower uterine segment cesarean section scar. Ultrasound Obstet Gynecol. 21 (3), 220-227 (2003).
  16. Fitzpatrick, K. E., et al. Incidence and risk factors for placenta accreta/increta/percreta in the UK: A national case-control study. PLoS One. 7 (12), e52893(2012).
  17. Jauniaux, E., Gronbeck, L., Bunce, C., Langhoff-Roos, J., Collins, S. L. Epidemiology of placenta previa accreta: A systematic review and meta-analysis. BMJ Open. 9 (11), e031193(2019).
  18. Silva, B., Viana Pinto, P., Costa, M. A. Cesarean scar pregnancy: A systematic review on expectant management. Eur J Obstet Gynecol Reprod Biol. 288, 36-43 (2023).
  19. Kaelin Agten, A., et al. First-trimester cesarean scar pregnancy: A comparative analysis of treatment options from the international registry. Am J Obstet Gynecol. 230 (6), 669.e1-669.e9 (2024).
  20. Gilman, A. R., Dewar, K. M., Rhone, S. A., Fluker, M. R. Intrauterine adhesions following miscarriage: Look and learn. J Obstet Gynaecol Can. 38 (5), 453-457 (2016).
  21. Smikle, C., Yarrarapu, S. N. S., Khetarpal, S. Statpearls. , Treasure Island (FL). (2024).
  22. Dreisler, E., Kjer, J. J. Asherman's syndrome: Current perspectives on diagnosis and management. Int J Womens Health. 11, 191-198 (2019).
  23. Hanuman, S., Pande, G., Nune, M. Current status and challenges in uterine myometrial tissue engineering. Bioengineered. 14 (1), 2251847(2023).
  24. Hanuman, S., B, H. K., Pai, K. S. R., Nune, M. Surface-conjugated galactose on electrospun polycaprolactone nanofibers: An innovative scaffold for uterine tissue engineering. ACS Omega. 9 (32), 34314-34328 (2024).
  25. Buhimschi, C. S., Zhao, G., Sora, N., Madri, J. A., Buhimschi, I. A. Myometrial wound healing post-cesarean delivery in the MRL/MPJ mouse model of uterine scarring. Am J Pathol. 177 (1), 197-207 (2010).
  26. Cowell, T. P. Implantation and development of mouse eggs transferred to the uteri of non-progestational mice. J Reprod Fertil. 19 (2), 239-245 (1969).
  27. Zhang, E. T., et al. Uterine injury during diestrus leads to placental and embryonic defects in future pregnancies in mice. Biol Reprod. , (2024).
  28. Zhang, E. T. Mouse surgical model of mechanical uterine injury and subsequent embryo defects. Curr Protoc. 4 (4), e1044(2024).
  29. Freedman, M. F., Schlaff, W. D. Avoiding Asherman's syndrome: Refining our approach to uterine evacuation. Fertil Steril. 116 (4), 961-962 (2021).
  30. Jensen, P. A., Stromme, W. B. Amenorrhea secondary to puerperal curettage (Asherman's syndrome). Am J Obstet Gynecol. 113 (2), 150-157 (1972).
  31. Li, Z., et al. Uterine scarring leads to adverse pregnant consequences by impairing the endometrium response to steroids. Endocrinology. 161 (11), bqaa174(2020).
  32. Burke, S. D., Zsengeller, Z. K., Karumanchi, S. A., Shainker, S. A. A mouse model of placenta accreta spectrum. Placenta. 99, 8-15 (2020).
  33. Anderson, J. M., Rodriguez, A., Chang, D. T. Foreign body reaction to biomaterials. Semin Immunol. 20 (2), 86-100 (2008).
  34. Carnicer-Lombarte, A., Chen, S. T., Malliaras, G. G., Barone, D. G. Foreign body reaction to implanted biomaterials and its impact in nerve neuroprosthetics. Front Bioeng Biotechnol. 9, 622524(2021).
  35. Klopfleisch, R., Jung, F. The pathology of the foreign body reaction against biomaterials. J Biomed Mater Res A. 105 (3), 927-940 (2017).
  36. Itoga, R., et al. Delayed onset of infectious foreign body granuloma for nonabsorbable sutures after open Achilles tendon repair: A case report. J Foot Ankle Surg. 60 (6), 1297-1300 (2021).
  37. Bowers, D., Mckenzie, D., Dutta, D., Wheeless, C. R., Cohen, W. R. Growth hormone treatment after cesarean delivery in rats increases the strength of the uterine scar. Am J Obstet Gynecol. 185 (3), 614-617 (2001).
  38. Kuramoto, G., et al. Human mesenchymal stem cell sheets improve uterine incision repair in a rodent hysterotomy model. Am J Perinatol. 39 (11), 1212-1222 (2022).
  39. Lin, N., et al. The effect of collagen-binding vascular endothelial growth factor on the remodeling of scarred rat uterus following full-thickness injury. Biomaterials. 33 (6), 1801-1807 (2012).
  40. Oh, S. S., Narver, H. L. Mouse and rat anesthesia and analgesia. Curr Protoc. 4 (2), e995(2024).
  41. Turner, P. V., Brabb, T., Pekow, C., Vasbinder, M. A. Administration of substances to laboratory animals: Routes of administration and factors to consider. J Am Assoc Lab Anim Sci. 50 (5), 600-613 (2011).
  42. Navarro, K. L., et al. Mouse anesthesia: The art and science. Ilar Journal. 62 (1-2), 238-273 (2021).
  43. Hoogstraten-Miller, S. L., Brown, P. A. Techniques in aseptic rodent surgery. Curr Protoc Immunol. Chapter 1 (Unit 1), 12-14 (2008).
  44. Stewart, K., Schroeder, V. A. Rodent identification I. J Vis Exp. , e10189(2023).
  45. Langford, D. J., et al. Coding of facial expressions of pain in the laboratory mouse. Nat Methods. 7 (6), 447-449 (2010).
  46. Ullman-Cullere, M. H., Foltz, C. J. Body condition scoring: A rapid and accurate method for assessing health status in mice. Lab Anim Sci. 49 (3), 319-323 (1999).
  47. Stewart, K., Schroeder, V. A. Fundamentals of breeding and weaning. J Vis Exp. , e10293(2023).
  48. Behringer, R., Gertsenstein, M., Nagy, K. V., Nagy, A. Selecting female mice in estrus and checking plugs. Cold Spring Harb Protoc. 2016 (8), prot092387(2016).
  49. Phoon, C. K. L., Ren, M. Is my mouse pregnant? High-frequency ultrasound assessment. J Vis Exp. (169), e61893(2021).
  50. Greco, A., et al. High-frequency ultrasound for in vivo pregnancy diagnosis and staging of placental and fetal development in mice. PLoS One. 8 (10), e77205(2013).
  51. Heyne, G. W., et al. A simple and reliable method for early pregnancy detection in inbred mice. J Am Assoc Lab Anim Sci. 54 (4), 368-371 (2015).
  52. Boerma, T., Ronsmans, C. Global epidemiology of use of and disparities in cesarean sections - Authors' reply. Lancet. 394 (10192), 25(2019).
  53. Alawadhi, F., Du, H., Cakmak, H., Taylor, H. S. Bone marrow-derived stem cell (BMDSC) transplantation improves fertility in a murine model of Asherman's syndrome. PLoS One. 9 (5), e96662(2014).
  54. Park, M., et al. Perivascular stem cell-derived cyclophilin A improves uterine environment with Asherman's syndrome via hif1alpha-dependent angiogenesis. Mol Ther. 28 (8), 1818-1832 (2020).
  55. Kim, J. H., et al. Intrauterine infusion of human platelet-rich plasma improves endometrial regeneration and pregnancy outcomes in a murine model of Asherman's syndrome. Front Physiol. 11, 105(2020).
  56. Feng, Q., et al. Establishment of an animal model of intrauterine adhesions after surgical abortion and curettage in pregnant rats. Ann Transl Med. 8 (4), 56(2020).
  57. Mansouri-Kivaj, N., et al. Homogenous subpopulation of human mesenchymal stem cells and their extracellular vesicles restore function of endometrium in an experimental rat model of Asherman syndrome. Stem Cell Res Ther. 14 (1), 61(2023).
  58. Gan, L., et al. Human amniotic mesenchymal stromal cell transplantation improves endometrial regeneration in rodent models of intrauterine adhesions. Cytotherapy. 19 (5), 603-616 (2017).

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Cesarean SectionPlacenta AccretaPlacenta PreviaUterine Wound HealingEmbryo ImplantationSurgical ProcedureUterine Scarring
Mouse Model of Surgical Uterine Injury and Subsequent Pregnancy Outcomes
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