Establishment of an Experimental Mouse Model of Endometrioma to Study its Related Infertility

Summary

This study introduces a novel mouse model specifically designed to simulate endometrioma, a clinically significant subtype of endometriosis. By utilizing C57BL/6J mice, the model aims to elucidate the pathophysiological mechanisms underlying endometrioma-related infertility, offering a refined tool to bridge the current knowledge gap in reproductive medicine.

Abstract

Endometrioma (OMA), a subtype of endometriosis characterized by the formation of endometriotic cysts in the ovaries, affects 17-44% of individuals diagnosed with endometriosis. Women with OMA often experience compromised fertility, yet the exact mechanisms underlying OMA-associated infertility remain unclear. Notably, existing animal models simulate superficial peritoneal endometriosis (SUP) and deep infiltrating endometriosis (DIE), leaving a notable gap in research focused on OMA. In response to the gap of knowledge, this paper introduces a pioneering OMA-simulating mouse model and provides a comprehensive description of the techniques and procedures employed in the model. With a high success rate of 83% and ovarian lesion specificity, this model holds significant promise for advancing our understanding of OMA, particularly in the context of infertility. It offers a valuable platform for conducting targeted research into OMA-associated fertility challenges, potentially paving the way for improved diagnostic and therapeutic strategies in the field of reproductive medicine.

Introduction

Endometrioma (OMA) is the most predominant subtype of endometriosis, observed in approximately 17-44% of individuals diagnosed with endometriosis^1,2. It is characterized by the formation of endometriotic cysts within the ovaries. These cysts, colloquially termed "chocolate cysts", derive their name from their distinctive brown, tar-like consistency³. In addition to OMA, endometriosis can also present as superficial peritoneal endometriosis (SUP) and deep infiltrating endometriosis (DIE). SUP refers to the lesions on the peritoneal lining, whereas DIE refers to lesions penetrating more than 5 mm underneath the peritoneal surface⁴. The heterogeneity in lesion location, appearance, and depth among these endometriosis subtypes results in diverse clinical symptoms and variable disease severity^5,6. OMA is particularly associated with more severe endometriosis stages and has been implicated in infertility, pelvic adhesions, and an increased ovarian cancer risk^1,7,8,9.

The association of endometriosis, especially OMA, with infertility is a clinical issue of grave concern. Endometriosis is present in 25-50% of infertile women, with 30-50% of women diagnosed with endometriosis experiencing infertility^{10,11,12,13,14}. While the exact OMA-associated infertility mechanisms remain elusive, some hypotheses have been raised. One suggests that endometriosis leads to a chronic inflammatory state, which can disrupt normal ovarian function and impair oocyte quality^15,16. Another proposes abnormal iron overload in the ovarian follicular fluid, which is believed to be associated with oocyte maturation disorder¹⁴. Other studies address the disruptions in hormonal regulation¹⁷, oocyte quality¹⁸, and embryo development¹⁹.

Historically, rodent models have played a crucial role in deepening our understanding of endometriosis, particularly in studying its pathology, impact on fertility, and mechanisms of pain^20,21,22,23. Rodents, particularly rats and mice, have been extensively utilized as animal models in exploring the cause-and-effect relationships, underlying mechanisms, potential diagnostic techniques, and therapeutic interventions for this disease^22,24,25,26. It is important to recognize that all animal models have their specific uses and limitations. The choice of a particular model depends on the specific outcome or aspect being measured or studied²⁷. For instance, a rat model demonstrated that stress could exacerbate endometriosis manifestations and influence inflammatory parameters, providing insights into the disease's potential triggers²⁸.

In the context of endometriosis research, there are various rodent models employed. One commonly used approach is the homologous model, which involves the transplantation of rodent uterine tissue into the same species' peritoneal cavity or mesenteric vessels²⁹. This model offers advantages in terms of immunocompetence and suitability for long-term studies³⁰. However, limitations arise due to the partial disparity between the implanted ectopic mouse uterine tissue and the characteristics of human endometriotic lesions³¹. Another approach is the heterologous model, where a human endometrial biopsy is implanted into an immunosuppressed mouse³². This model allows the use of human ectopic endometrium as a donor tissue for lesion development, providing constructive validity³². However, it relies on immunosuppressed mice as recipients, which hinders a comprehensive assessment of the immune responses involved in the etiology of the disorder³³. Nevertheless, it is important to acknowledge that the existing models primarily focus on mirroring the generalized condition SUP, inadequately capturing the unique characteristics of OMA and DIE³⁴. Currently, there is a paucity of specific models available for studying DIE, with only a few models existing, including a recent rodent model developed by Yan et al.³⁵. This scarcity underscores the need for further research and the development of models that accurately capture the specific characteristics of DIE. Additionally, our understanding of OMA, especially its nuanced association with fertility, also remains limited^26,36.

Addressing the existing challenges, this paper presents a novel experimental homologous mouse model designed to specifically simulate OMA. It is aimed at providing unique insights into the pathological mechanisms underlying OMA-related infertility, thereby bridging the knowledge gap in this crucial area of reproductive medicine. In brief, C57BL/6J mice were used for their human-like reproductive traits. Following estrus cycle synchronization, uterine tissues from donor mice were minced and transplanted into the recipient mice's ovarian bursa at a 1:2 ratio. After a 4-week interval, both morphological and histological examinations of the ovarian lesions were conducted to validate OMA presence and evaluate any associated follicular atresia. With an 83% successful rate, this model offers researchers a reliable platform for OMA studies, emphasizing lesion development specifically in the ovaries for focused investigations.

Protocol

All experimental procedures conducted in this study were approved and regulated under protocol number 21-203-MIS_[C] by the Ethical Committee of the Chinese University of Hong Kong.

1. Animal acclimatization and selection

1. Mice preparation
   1. Obtain 18 C57BL/6J mice (primiparous females, 8-9 weeks old: 6 donors, 12 recipients) from a reputable supplier. Verify the health certificates.
   2. Maintain all mice in a controlled housing environment: 22-24 °C, humidity levels between 40% and 60%, and with a 12 h light/dark cycle. Use HEPA-filtered air, if available.
   3. Allow 72 h acclimation for the mice with continuous access to food and water, minimizing human interaction to reduce stress.
2. Estrous cycle synchronization
   1. Collect bedding containing male pheromones; introduce it to female cages for 48 h.
3. Vaginal cytology
   1. Perform vaginal cytology the following morning.
      NOTE: It is necessary to perform a smear at a consistent time of day, typically 9:00 AM.
      1. Prepare double-headed cotton swabs, autoclaved PBS, a 35 mm Petri dish, and glass slides in advance. Dispense 10 mL of PBS in a Petri dish. Submerge one end of the cotton swab in the PBS to allow it to absorb the liquid.
      2. Gently take out one mouse from its cage and place it on the lid of an empty cage.
      3. Grasp the tail using the thumb and index finger of one hand, and with the other hand take the moistened cotton swab and delicately insert it into the mouse's vagina. Carefully insert the tip of the swab stick to a depth of approximately 1.0 cm into the mouse's vagina. Rotate the swab gently, while maintaining an angle of about 45° to the long axis of the animal's body.
         NOTE: Rotating the swab slowly during insertion can facilitate a smoother procedure and minimize the potential stimulation to the cervix.
      4. Gently remove cells from the vaginal lumen and walls by continuously rotating the swab, and then carefully withdraw the swab.
         NOTE: During this process, ensure that the swab is continuously rotated while being cautious not to come into contact with the surrounding hairs. Afterward, carefully withdraw the swab to avoid any potential contamination.
      5. Transfer the collected cells by gently rolling the swab tip onto a clean, prelabeled glass slide.
         NOTE: The swab stick should be discarded after transferring cells to slides, and a new one should be used for each animal.
      6. Inspect the slides under a microscope to detect cornified epithelial cells. Look for the uniform presence of cornified epithelial cells in vaginal cytology, which is indicative of the estrus stage. Capture photographs and record the observations.
         NOTE: With the appropriate expertise, staining of vaginal smears is not necessary. However, if an experimenter cannot "read" the smears without them being stained, then it is necessary.

2. Endometrioma model establishment

NOTE: Only select mice at the estrus stage as donor and recipient mice in the model establishment. When performing experiments on one mouse, ensure it is out of the sight of others and that it is not reintroduced to the company of other animals until completely recovered. The diagram of model generation can be found in Figure 1.

1. Donor tissue preparation
   1. Sedate the donor mice with a combination of ketamine (75 mg/kg) and xylazine (10 mg/kg). Euthanize the donor mice by cervical translocation under anesthesia.
   2. Disinfect the abdominal area with a 70% ethanol swab.
   3. Use sterile surgical scissors to create an incision (1 cm) along the midline of the peritoneum to expose the muscular layer. Choose another pair of surgical scissors to cut the muscle layer and reveal the pelvic cavity.
   4. Close the tips of the forceps and gently lift the intestines using the blunt central portion.
   5. Locate the uterus and ovarian tissues. Conduct a sterile dissection of the whole uterine horn, including both uterine horns, making sure that the approximate size of the piece being removed is ~1.5 cm in length. Place the excised tissues in a petri dish containing PBS and remove any excess fat tissues.
   6. Minutely dissect the tissues into 1 mm³ pieces using a sterilized scalpel. Keep the tissues moist by periodically adding PBS.
2. Transplantation procedure
   1. Anesthetize the recipient mice with a mixture of ketamine (75 mg/kg) and xylazine (10 mg/kg).
   2. Apply ophthalmic ointment to prevent eye dryness during the surgery. Place the mice in a supine position on a sterile surgical board.
   3. Determine the surgical site by gently manipulating the hind legs of the mouse, with a bony prominence palpable under the skin. This point of prominence, clearly visible through the skin, serves as a reliable anatomical marker for the ovaries of the recipient mouse during the surgical procedure.
   4. Disinfect the surgical area with a 70% ethanol swab. Create a precise lateral incision (3-5 mm) on the predetermined surgical site.
   5. Stabilize the ovary with non-dominant hand forceps. Gently inject 100 µL of PBS into the ovarian bursa, observing the slight separation.
   6. Create a tiny slit in the bursa. Transplant the uterine tissue fragments (1 mm³) using micro forceps.
      NOTE: Avoid overstuffing to prevent pressure on surrounding tissues.
   7. For sham surgery, replicate all steps except step 2.2.6.
3. Postoperative care
   1. Suture the muscle layers with absorbable sutures (i.e., 5-0 vicryl) and skin with non-absorbable sutures (i.e., 5-0 nylon).
   2. Place a mouse on a heating pad set to 37 °C until full consciousness returns. Monitor the breathing and wait for the extremities to turn pink.
   3. Administer buprenorphine (0.1 mg/kg, subcutaneously) for pain every 12 h for 48 h. Check for signs of pain, infection, or distress every 8 h for the first 24 h and then 3x daily for the next 3 days.
      NOTE: The animal is not left unattended until it has regained enough consciousness to sustain sternal recumbency.

3. Model validation

1. Gross validation
   1. After 4 weeks, anesthetize the recipient mice using a combination of ketamine (75 mg/kg) and xylazine (10 mg/kg). Then, under deep anesthesia, euthanize the mice by performing cervical translocation.
   2. Perform a midline incision and expose the abdominal cavity for organ inspection.
   3. Extract the ovaries with visible lesions and take photographs.
   4. Thoroughly inspect for extraneous endometriotic lesions.
2. Histological validation
   NOTE: Due to the presence of potentially irritating odoriferous solvents in the histological experiments, it should be conducted in a fume hood.
   1. Fix the ovaries together with lesions in 10% neutral-buffered formalin.
   2. Process the tissues following the same procedures as described by Adeniran et al.³⁷, including tissue allocation, fixation, and embedding.
   3. Serially section tissues at 4-5 µm thickness, float in a water bath at 45 °C, and mount on glass slides. Clearly label the slides with mouse number, ovary side, and section number. Dry overnight at 37 °C.
   4. PAS-hematoxylin staining
      1. Deparaffinization: Place the slides in xylene or a xylene substitute to remove the paraffin.
         NOTE: You may need to perform this step a couple of times.
      2. Rehydration: Gradually rehydrate the tissue sections by placing the slides in a series of decreasing ethanol concentrations (100%, 95%, 80%, 70%).
      3. Periodic acid treatment: Cover the tissue sections with a 1% periodic acid solution and allow them to sit for 5-10 min at room temperature.
         NOTE: This step oxidizes the tissue's glycol groups to aldehyde groups, allowing for reaction with the Schiff reagent and the formation of a purple-pink stain to indicate the ovarian structures.
      4. Rinse the slides thoroughly with distilled water to remove any residual periodic acid.
      5. Stain the tissue sections with a Schiff's solution for 15 min.
      6. Rinse the slides first in hot running tap water to remove excess stain, then in distilled water.
      7. Counterstain the sections with 1 g/L of Meyer's hematoxylin for 2-3 min.
      8. Rinse slides in running tap water for 2-3 min.
      9. Apply Bluing Reagent for 30 s.
      10. Rinse slides in distilled water.
      11. Gradually dehydrate the sections by immersing them in increasing ethanol concentrations (70%, 80%, 95%, and 100%).
      12. Submerge the sections in a clearing agent (e.g., xylene) to make them transparent.
      13. Apply a mounting medium to the sections on the slides and gently place a coverslip over the sections, ensuring there are no air bubbles. Allow the slides to air-dry in the hood to facilitate the solidification of the mounting medium.
   5. Examine under a light microscope. Document the presence of endometrial glands, stroma, and hemorrhagic cysts confirming endometriosis in the ovary.

Results

The successful establishment of OMA
Of the 12 mice subjected to the transplantation protocol, 10 exhibited characteristic OMA lesions, both at a gross anatomical and histological level, translating to a success rate of 83%. Gross examination revealed fluid-filled lesions adherent to the ovaries, reminiscent of clinical OMA presentations (Figure 2). Histopathological scrutiny, as illustrated in Figure 3, confirmed the ectopic endometrial ti...

Discussion

The prevalence of OMA in the global female population underscores a critical health concern³⁸. Beyond the general symptoms of endometriosis, OMA brings additional challenges for fertility, including severe pelvic pain, potential for ovarian torsion, and other notable implications^39,40. While the understanding of OMA pathogenesis and progression is largely shrouded in mystery, the knowledge vacuum not only impedes the formulation of targete...

Materials

Name	Company	Catalog Number	Comments
1 cc syringe with 25 G needle	BD Biosciences	301320
10 mm Tissue culture dish	FALCON	353001
10% Buffered formalin	Fisher Scientific	SF100-4
10x phosphate buffered saline (PBS) buffer	Fisher Scientific	AM9625
30 G needle	BD Biosciences	305107
5-0 black braided silk non-absorbable suture	Ethicon Inc.	W500H
70% Ethanol
Adhesive microscope slides	Marienfeld	810401
Buprenorphine Injection	Med-Vet International	RXBUPRENOR5
Double-headed cotton swab	SANYO Co., LTD.	HUBY-340
Fine forceps	Fine Science Tools	11254-20
Ketamine	Alfasan International b.v	2203095-08
Microtubes	Corning	MCT-150-C
Needle holder	Excelta Corporation	2827-NH-35-SE-ND
Ophthalmic ointment	Major Pharmaceuticals	10033691
Periodic Acid Schiff (PAS) Stain Kit	Abcam	Ab150680
Powder free sterile gloves	Fisher Scientific	19020558
Processing/embedding cassettes	Fisher Scientific	15-197-700A
Size 3 scalpel	Fisher Scientific	22-079-657
Small serrated semi-curved forceps	Roboz Surgical Instruments Co.	RS-5135
Small surgical scissors	Roboz Surgical Instruments Co.	RS-5850
Standard light microscope			For evaluating vaginal cytology smears and histological analysis.
Steel scalpel blades #10	B.Braun	BB510
Sterile gauze	MEDICOM	HMWS 077
Sterilized pyrex glass Petri dishes	Corning	70160-101
Thermoregulated electric pad
VICRYL RAPIDE (polyglactin 910) absorbable Suture	Ethicon Inc.	W9918
Xylazine	Alfasan International b.v	2110333-10