Protocol for ovariectomy and estradiol replacement in mice

Summary

Ovariectomy, involving the surgical removal of ovaries, and estradiol replacement facilitate the understanding of sexual dimorphism-related physiological changes, encompassing reproductive biology, metabolism, and hormone-related diseases. In this study, we present a protocol for conducting ovariectomy and estradiol replacement in mice. We describe steps for performing sham and ovariectomy operations, outline preoperative preparations, and provide details on postoperative care, including analgesia administration and the removal of surgical clips. Additionally, we elaborate on the procedures for performing vehicle and estradiol injections.

For complete details on the use and execution of this protocol, please refer to Luengo-Mateos et al.¹

Graphical abstract

Highlights

• •A guide with detailed procedural steps for performing ovariectomy in mice
• •Instructions covering pre-ovariectomy preparations and postoperative care
• •Steps and considerations for administering estradiol to ovariectomized mice

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Ovariectomy, involving the surgical removal of ovaries, and estradiol replacement facilitate the understanding of sexual dimorphism-related physiological changes, encompassing reproductive biology, metabolism, and hormone-related diseases. In this study, we present a protocol for conducting ovariectomy and estradiol replacement in mice. We describe steps for performing sham and ovariectomy operations, outline preoperative preparations, and provide details on postoperative care, including analgesia administration and the removal of surgical clips. Additionally, we elaborate on the procedures for performing vehicle and estradiol injections.

Before you begin

Menopause, characterized by declining ovarian function and lower estrogen levels, significantly impacts health, leading to physiological shifts in metabolism and immune responses.² Post-menopausal individuals face elevated mortality risks, especially from conditions like heart disease, breast cancer, and endometrial cancer.² These changes in mortality patterns are often attributed to the loss of protective effects from reproductive hormones, particularly estrogens. Estimations indicate that by 2030, the number of menopausal individuals will reach 1.2 billion, with 47 million people transitioning to post-menopause each year until then.³ The global aging population is on the rise, and projections suggest a significant increase, underscoring the complexity and incomplete understanding of menopausal physiology, emphasizing the urgent need to enhance research efforts. Grasping these changes is crucial for addressing the health challenges associated with this life stage.

In scientific research, ovariectomy plays a pivotal role in investigating sexual dimorphism-related physiological changes and the broader field of reproductive biology. Researchers employ this technique to explore its physiological impact and uncover the underlying mechanisms contributing to sex-specific differences in metabolism, hormone-related diseases, and energy homeostasis. As the importance of ovarian hormones becomes more evident, the ovariectomy protocol remains an indispensable tool for achieving critical breakthroughs in these vital areas of study.

The primary objective of this protocol is to provide a comprehensive guideline for performing ovariectomy in mice, covering the use of the isoflurane anesthesia system, postoperative care procedures, and methods for evaluating the surgery’s success. Ovariectomy can be performed using various approaches, such as a single surgical incision, double dorsolateral incisions, or a single ventral transverse incision in the middle part of the abdomen. Our research group’s experience indicates that double dorsolateral incisions offer advantages, including enhanced access to the ovaries and quicker postoperative recovery. This method provides a detailed description of the ovariectomy surgical procedure, with a particular focus on the double dorsolateral incision method. Additionally, we outline the steps for conducting the estradiol replacement experiment, where estradiol is administered to ovariectomized mice to restore their endocrine function. This experiment enables scientists to investigate the impact of estradiol action precisely and systematically in OVX animals.

Our goal is to enhance the accuracy and consistency of ovariectomy procedures, ensuring the welfare of animals and adhering to ethical and safety principles by having qualified and trained professionals conduct all procedures.

Institutional permissions

All animal care procedures strictly adhered to the guidelines established by the institutional animal care committee. These procedures underwent comprehensive review and received approval from the University of Santiago de Compostela Ethics Committee, in accordance with the European Union’s regulatory framework governing the use of experimental animals (Project ID 15012/2021/011).

For a brief overview, the mice were housed with unrestricted access to both food and water and were kept in a controlled environment. They were maintained under a 12-h light-dark cycle (from 8 a.m. to 8 p.m.) in a room with controlled humidity and temperature (21°C) within the University of Santiago de Compostela’s animal facility. We strongly recommend obtaining the necessary permissions from the relevant institutions before commencing any research procedures.

Ovariectomy: Preoperative preparation

Timing: 2 h

This section outlines the essential materials, equipment, and solutions that must be prepared before the surgery.

• 1.Sterilization of surgical equipment: Surgical instruments are placed in a sterile pack and subjected to steam under pressure. The use of a sterilization indicator, such as autoclave tape or a strip, is crucial to verify the sterilization process. Refer to Figure 1 for a depiction of the required instruments for the ovariectomy.

CRITICAL: Ensure meticulous sterilization of all surgical instruments to maintain aseptic conditions throughout the procedure. Proper sterilization is crucial to prevent infections and ensure the success of the surgical procedure. Autoclave sterilization is the most employed method for instrument sterilization.

• 2.
  Preparing anesthesia and analgesia solutions.

  • a.
    Prepare the solutions by determining the required concentration based on the weight and dosage appropriate for your specific animal model.
  • b.
    Dilute the drugs in an appropriate solvent, typically sterile saline, to achieve the desired concentration.
  • c.
    Calculate and prepare individual doses for each animal based on their weight and the predetermined dosage.
  • d.
    Ensure thorough mixing of the solution, and label it clearly with information including the concentration and expiration date.

    • i.
      Meloxicam for pre-operative analgesia.
    • ii.
      Ketamine/Xylazine solution for anesthesia.
    • iii.
      Meloxicam for routine post-operative analgesia. In cases where severe pain signs are observed, judiciously use buprenorphine to provide additional relief.

Note: It is of utmost importance to adhere to institutional guidelines, legal considerations, and safety protocols when handling and administering these solutions to guarantee the welfare of the animals involved in the study.

• 3.
  Isoflurane utilization with intraperitoneal anesthesia. To set up the veterinary anesthesia system for isoflurane administration, please follow these steps using Figure 2 as a reference:

  • a.
    Setting up the isoflurane flow: adjust the oxygen flow in the flowmeter to a range of 0.8–1 L/min for oxygen, ensuring that the valve is open.
  • b.
    Isoflurane absorption: activate the fluosorber to absorb isoflurane from the system.
  • c.
    Adjust the isoflurane concentrator: maintain the isoflurane concentrator between 1%‒2%. This device facilitates the evaporation of liquid isoflurane into the air for administration.
  • d.
    Open the valves in the Isoflurane nose cone to allow the controlled delivery of isoflurane to the animals.

Note: In this protocol, we combine isoflurane with intraperitoneal anesthesia to enhance analgesia and muscle relaxation.

• 4.Establishing a post-surgery area. Create a designated post-surgery area equipped with a heating pad, where mice can be placed until they fully regain consciousness.

Figure 1.

Surgical instruments used for ovariectomy, from left to right: surgical clip applier, surgical clip remover, fine curved scissors, fine tissue forceps, fine dressing forceps, needle holder, and scissors

Figure 2.

Veterinary isoflurane anesthesia system

(A–D) Anesthesia System Components: (A) Flowmeter. (B) Fluosorber. (C) Isoflurane vaporizer. (D) Isoflurane Cone. See also Figure 4.

Following these steps will effectively set up the isoflurane administration system and provide a safe and controlled environment for anesthesia and post-surgery care.

Estradiol replacement

Timing: 1 week

• 5.
  Animal Housing:

  • a.
    Individually house the mice to facilitate precise measurement of food intake.

    Note: This housing approach is specifically designed for scientific accuracy in food intake assessment and is not intended to disregard mice social nature.

  • b.
    To minimize stress and reduce variations in response due to environmental factors, allow mice to acclimatize to their housing conditions for at least 3 days before the experiment.
• 6.
  Estradiol solution preparation:

  • a.
    Vehicle solution: prepare 150 μL of sesame oil for each mouse.
  • b.
    Estradiol solution: Dissolve 1 μg of 17-β-estradiol-3-benzoate (E8515, Sigma-Aldrich) in 150 μL of sesame oil for each mouse.
  • c.
    To facilitate the dissolution of estradiol, heat it to 40° and let it stir for 12–16 h. Other dissolving methods might be applicable, but this is the one we have solid experience with.
  • d.
    Store the solution in a light-protected environment to prevent degradation.

Note: Store the estradiol solution at 20°C–22°C for the entire experiment. For long-term storage, use glass tubes, as steroids tend to adhere to the walls of plastic and polypropylene tubes.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Chemicals, peptides, and recombinant proteins
Buprenorphine (Bupaq)	Richter Pharma	Cat. No. 578816.6
Ketamine (Ketamidor)	Richter Pharma	Cat. No. 580393.7
Meloxicam (Metacam)	Boehringer Ingelheim	Cat. No. 52044186/006 52044186/006
Xylazine hydrochloride (Rompun)	Bayer	Cat. No. 572126.2
Isoflutek 1000 mg/g	Karizoo	Cat. No. 586259.0
b-Estradiol 3-benzoate	Sigma-Aldrich	Cat. No. E8515
Sesame oil	Sigma-Aldrich	Cat. No. S3547
Chlorhexidine digluconate 2%	Alvita	Cat. No. 180584.3
Saline vitulia	ERN	Cat. No. SF-VIT-500
Fast Panoptic kit	PanReac AppliChem	Cat. No. 254807
Experimental models: Organisms/strains
Mouse, C57BL/6J, adult, female	The Jackson Laboratory	N/A
Other
Puralube vet ophthalmic (eye) ointment	MWI Veterinary	Cat. No. 027505
Autoclip 9 mm applier	Stoelting	Cat. No. 59043
Autoclip remover	Stoelting	Cat. No. 59046
Micro-Adson forceps	Fine Science Tools	Cat. No. 11018-12
Strabismus scissors	Fine Science Tools	Cat. No. 14074-09
Student Halsey needle holder	Fine Science Tools	Cat. No. 91201-13
Tissue forceps - 1 × 2 teeth	Fine Science Tools	Cat. No. 11021-12
Surgical scissors	Fine Science Tools	Cat. No. 14002-13
Autoclips	Mikron Precision, Inc.	Cat. No. 205016
Glass bead sterilizer (Steri 250)	Sigma-Aldrich	Cat. No. Z378569
Autoclave tape	Dental Iberica	Cat No. OM152
Heat pad	Pekatherm	Cat. No. USD20TD
Electric razor (Baby Beast trimmer)	Skull Shaver	Cat. No. BBTEU
Sterile drape	Glad	Cat. No. 3919849

Materials and equipment

Prepare the anesthesia, analgesia, and estradiol solutions as described below:

Meloxicam: Prepare a 5 mg/mL solution by mixing 1 mL of Meloxicam with 9 mL of saline solution. Administer 0.1 mL of the prepared solution subcutaneously per 10 g of the animal’s weight (dose = 5 mg/kg).

Ketamine/Xylazine: Prepare a 10 mg/mL Ketamine and 2 mg/mL Xylazine solution by mixing 1 mL of Ketamine (100 mg/mL), 0.5 mL of Xylazine (20 mg/mL), and 8.5 mL of saline solution. Administer 0.05 mL of the prepared solution intraperitoneally per 10 g of the animal’s weight (dose = Ketamine: 50 mg/kg, Xylazine: 5 mg/kg).

Buprenorphine: Prepare a 0.3 mg/mL solution by mixing 1 mL of Buprenorphine with 9 mL of saline solution. Administer 0.05–0.1 mL of the prepared solution subcutaneously per 30 g of the animal’s weight (dose = 0.05–0.1 mg/kg).

17-β-estradiol-3-benzoate: Prepare a 6.66 ng/mL solution by mixing 100 μg of 17-β-estradiol-3-benzoate in 15 mL of sesame oil for each mouse (heat it to 40°C and let it stir for 12–16 h). Administer 150 μL of the prepared solution subcutaneously per animal (dose = 150 μL/mice/day).

Step-by-step method details

Ovariectomy procedure

Timing: 1 week

The following steps are crucial to ensure a successful ovariectomy procedure, emphasizing precision and caution to minimize the risk of complications while safeguarding the well-being of the animals.

Please refer to Figure 3 for visual guidance, which includes photographs depicting the most important steps of the OVX. Additionally, Figure 4 is a diagram illustrating the sequential steps of the ovariectomy procedure.

• 1.
  Preoperative Preparation: 15–20 min.

  • a.
    Administer the Meloxicam solution subcutaneously to provide analgesia.
  • b.
    Administer the Ketamine/Xylazine solution intraperitoneally to induce anesthesia. Ensure that the mouse is anesthetized within 10 min.
  • c.
    Apply Pura lube ointment to the eyes to prevent dryness.
  • d.
    Gently press the footpad of the animal to ensure desensitization.
  • e.
    Carefully shave the right and left flanks of the mice using an electric razor.
  • f.
    Position each mouse in the anesthesia isoflurane mask, lying on its left flank over a warm electric blanket (Figure 4).
  • g.
    Confirm deep anesthesia by gently pressing the footpad.
  • h.
    Thoroughly disinfect the flank using a 2% solution of chlorhexidine digluconate.
  • i.
    Place a sterile drape with cutout over the mouse, only exposing the surgical area (Figure 3A).
• 2.
  Ovariectomy: 20 min.

  • a.
    Create a small incision on the disinfected flank, ensuring that you cut through the skin, subcutaneous fat, and muscle layer (3–5 mm).

    CRITICAL: Exercise extreme caution to avoid accidentally catching the intestine while cutting through the muscle layer.

  • b.
    Ovary removal:

    • i.
      Gently lift the fat pad outwards using forceps to exteriorize the ovary and uterus through the incision.
    • ii.
      To prevent bleeding, secure the distal part of the uterine horn using a simple surgical suture (Figures 3A and 3B).
    • iii.
      Perform the ovary removal by dissecting the tissue between the suture and the ovary. Refer to Figures 3 and 4 for a visual guide.

      Note: Ensure that the suture is not placed too close to the ovary to prevent it from coming undone during the ovary dissection.

  • c.
    Closure:

    • i.
      Carefully place the uterine horn back into the abdominal cavity.
    • ii.
      Suture the muscle layer using a continuous technique with absorbable surgical sutures to prevent herniation or complications (Figure 3C).
    • iii.
      Use surgical clips to carefully close the skin wound, ensuring no sutures are left to minimize infection or irritation.

      Note: Proper closure is crucial for the success of the ovariectomy procedure and for the postoperative recovery of the animals. Thoroughness and meticulousness during this phase promote the well-being of the mice and minimize potential complications.

  • d.
    Repeat the same procedure on the right flank.
  • e.
    Disinfect the instruments between each animal by introducing them into a glass bead sterilizer (hot glass bead sterilizer).

    CRITICAL: Keep the surgery within a maximum limit of 25 minutes to prevent premature waking and excessive loss of body temperature. Strict timing ensures animal well-being and successful recovery.

• 3.
  Postoperative Care: 7–10 days.

  • a.
    Administer subcutaneous buprenorphine for postoperative pain management.
  • b.
    To prevent mouse dehydration during surgery, inject pre-warmed (37°C) 0.9% saline subcutaneously.

    Note: The recommended administration volume for mice is 20 mL/kg.

  • c.
    Place the mouse on a moderately warm electric blanket until fully conscious.

    Note: Proper postoperative care is crucial for ensuring the well-being and recovery of the animals, minimizing pain and complications to ensure comfort and health during the recovery period. It is essential to provide a comfortable and monitored environment during recovery, and to monitor the animals until they are fully recovered before returning them to the housing room.

  • d.
    After 24 h post-surgery, provide intraperitoneal meloxicam to deliver pain relief and support the recovery process. Repeat for 2–3 days.
  • e.
    7–10 days post-surgery, use a specialized clip remover to gently eliminate surgical clips, preventing discomfort or complications during healing.
  • f.
    Confirm the efficacy of the surgery by assessing serum estradiol levels using commercial estradiol immunoassays, as described in Haisenleder et al., 2011.⁴

    Note: The concentration is expected to decrease to 20–30% of the baseline after 2 weeks post-ovariectomy.⁵

Figure 3.

Ovary removal

(A and B) (A) The blue line represents the placement of the surgical suture (B) within the oviduct. The purple, discontinuous lines indicate the precise locations where the ovary should be cut during the removal procedure.

(C) Depicts the continuous muscle suture. See also Figure 4.

Figure 4.

Diagram showing the steps of the ovariectomy procedure

The first step (1) of the diagram illustrates the isoflurane anesthesia system and the positioning of the mice on a warming blanket for anesthesia induction. In the second step (2), the diagram highlights the anatomical location of the mouse ovaries, which are situated at the posterolateral poles of the kidneys, with each ovary attached by the mesovarium to the dorsal body wall of the abdominal cavity. The diagram also indicates the placement of a single suture within the oviduct during the ovariectomy procedure. The third step of the diagram (3) depicts the mice after the ovariectomy procedure, showing the presence of wound clips. Created in BioRender.com.

Sham procedure

Timing: 1 week

The sham procedure establishes a corresponding control group, mimicking the surgical steps of the experimental group without actual ovary removal.

• 4.Preoperative preparation: follow the previously described preoperative steps, which include anesthesia, shaving, analgesia administration, hydration, anesthesia mask placement, flank disinfection and the delineation of the surgical area.
• 5.Ovariectomy step (excluded): skip the ovary removal step. Instead, gently expose the ovaries and uterine horns outside of the abdominal cavity without proceeding to removal.
• 6.Return ovary and uterine horn: after exposing the ovaries and uterine horns, return them to the abdominal cavity.
• 7.Closure: follow the closure steps as previously outlined, which include suturing the muscle and closing the skin wound.

Estradiol treatment

Timing: 1 week

Following ovariectomy, the decline in ovary-derived hormones, particularly estradiol, prompts the estradiol replacement experiment in OVX animals. This experiment allows for the specific analysis of estradiol’s endocrine effects on reproductive, bone density, and metabolism studies.^1,6,7,8,9

Before initiating estradiol treatment, establish baseline days by performing daily subcutaneous injections of 150 μL of sesame oil to the mice over 3 days. This helps account for potential stress caused by injections.

• 8.Vehicle injections: using sterile syringes and needles, perform daily subcutaneous injections of 150 μL of sesame oil to the mice over the baseline days for 3 days.
• 9.Estradiol Injections: administer daily subcutaneous injections of the estradiol solution (1 μg of 17-β-estradiol-3-benzoate in 150 μL of sesame oil for each mouse over the next five days.

Expected outcomes

Based on our previous experience, we can anticipate specific outcomes, particularly in the context of energy balance physiology. Reduced estradiol levels resulting from ovarian insufficiency, whether due to physiological (menopause) or surgical (ovariectomy) causes, are associated with hyperphagia, decreased energy expenditure, and weight gain^1,8,9 In such scenarios, estradiol replacement can effectively prevent or reverse ovariectomy-induced obesity by reducing energy intake and increasing energy expenditure. The method detailed here has been standardized to ensure consistent and optimal results. Thus, we next outline the anticipated outcomes of the ovariectomy procedure:

Immediate recovery: mice should fully recover within 1 h of surgery and regain their ability to walk.

Short-term recovery: complete recovery, including resumption of normal activities, should be observed during the first week post-surgery.

Body weight: After approximately one week post-surgery, mice are expected to regain their baseline body weight.^1,10

Weight changes: Ovariectomized control mice are likely to experience weight gain due to the absence of estrogens, while sham-operated mice should maintain their baseline body weight. It’s essential to note that an increase in body weight and hyperphagia is typically observed starting from the second or third week following the surgery. This is primarily due to the gradual washout of ovarian hormones, a process that takes at least two weeks. Therefore, all experimental tests on OVX mice should be conducted at least two weeks after surgery to ensure the complete elimination of ovarian hormones.

Estradiol replacement: Following estradiol replacement, ovariectomized females are expected to reduce their food intake. After five days of estradiol replacement, ovariectomized females should exhibit reduced food intake compared to ovariectomized controls injected with the vehicle.

Limitations

While our protocol is optimized to ensure consistent and reliable results in our research, it’s essential to recognize several limitations associated with the OVX procedure and estrogen replacement.

Ovariectomy is an invasive surgical procedure: it raises ethical concerns regarding animal welfare. Therefore, strict adherence to ethical guidelines and regulations is imperative to safeguard the well-being of the animals involved in the study and address these ethical considerations adequately.

Potential induction of stress responses: the surgical procedure itself has the potential to induce stress responses in animals, which may influence study outcomes and introduce confounding variables. Therefore, it is crucial to diligently monitor and account for stress-related effects in experimental design and data analysis.

Variability in individual responses: The unique responses of individual animals to the OVX procedure can introduce experimental variability. This variability should be taken into consideration when interpreting results and might necessitate larger sample sizes to ensure statistical robustness.

Challenges in estradiol delivery: When considering various methods for estradiol delivery in mice, subcutaneous options such as subcutaneous injections, implants of estradiol pellets, or silastic tubes filled with estradiol powder have their limitations.¹¹ With subcutaneous injections, two limitations should be considered: a) the injected volume accumulates in a subcutaneous 'pocket,' affecting pharmacokinetics and preventing a drastic washout of the treatment. In fact, these pockets are still present weeks after the injection. b) Daily injections for long periods of time should be avoided to preserve animal well-being. Regarding subcutaneous implants, none of the procedures ensures the controlled release of estradiol. Typically, they lead to an initial high hormone delivery shortly after implantation, followed by an acute drop in release. In contrast, implants of osmotic mini pumps offer a solution to this issue, as the pores in the filters regulate the amount of solution delivered. However, the dimensions of the osmotic pump and the volume of the solution loaded are limited due to the small size of the mice. Additionally, surgical removal of the pump is normally necessary after 4 weeks, requiring a replacement or the finalization of the treatment.¹¹

Finally, for all these estradiol deliveries, hormone administration does not replicate the normal cyclic fluctuations of estradiol in females. Therefore, caution should be taken when interpreting the results.

Impact of subcutaneous injections: during estrogen replacement, it’s important to note that subcutaneous injections can have localized effects on tissue and blood flow, potentially influencing the absorption and distribution of estradiol. Furthermore, daily subcutaneous injections of estradiol may not precisely replicate physiological conditions, where estradiol levels oscillate along the estrous cycle, potentially leading to discrepancies in hormone levels.

Lastly, the selected dose (i.e., 1 μg/mice/day of 17-β-estradiol-3-benzoate) has been used by us and others^8,12 to study estradiol physiological actions. However, if the treatment is prolonged for several weeks, it may result in supraphysiological concentrations in plasma. If prolonged treatments are necessary for the experimental paradigm, researchers should assess potential undesired effects on estradiol-responsive tissues, such as uterus hypertrophy and hyperplasia.

Long-term health considerations: in long-term studies, the absence of ovarian hormones resulting from ovariectomy can lead to health issues in animals, potentially affecting the interpretation of results. Researchers must be mindful of potential long-term consequences and incorporate them into the study’s design.

Acknowledging and addressing these limitations is vital for the ethical and scientific rigor of research involving the OVX procedure and estrogen replacement.

Troubleshooting

Problem 1

Inadequate breathing in mice during anesthesia (related to steps 1f and 1g).

Potential solution

Inadequate breathing can result from excessive anesthesia. To address this issue, gradually reduce the isoflurane concentration until normal breathing is restored.

Problem 2

Hypothermia during and after the surgical procedure (related to step 1f).

Potential solution

To prevent hypothermia during or after the surgical procedure, follow these steps.

• •Maintain a warm environment for the mouse by placing it on a moderately warm electric blanket or a heating lamp.
• •Ensure the mouse is fully awake and alert before removing it from the warm environment.
• •Limit the surgery duration to avoid exceeding 25 min, thus minimizing body temperature loss.

Problem 3

Intestine injury during the surgery procedure (related to step 2a).

Potential solution

To prevent intestine injury, exercise extreme caution when making the incision on the flank. Ensure that you do not damage the intestine while cutting through the muscle layer. If there is a suspicion of intestinal damage, it is advisable to euthanize the mouse to prevent further suffering.

Problem 4

Bleeding during ovary removal (related to step 2b).

Potential solution

To address bleeding during ovary removal, it’s important to maintain a sufficient distance between the suture and the ovary to prevent it from getting unpicked when dissecting the ovary. Additionally, cauterizing the ovary is an alternative method that can be employed to manage bleeding during the procedure. If excessive bleeding occurs and cannot be controlled, it is advisable to sacrifice the mouse to minimize distress and harm. Proper technique and care in suturing are crucial to prevent complications during the procedure.

Problem 5

Bleeding and open wound during postoperative care (related to step 2c).

Potential solution

To address bleeding and open wounds during postoperative care.

• •When closing the skin, ensure that there is no suture outside of the wound, and that the clips are tightly secured.
• •If significant bleeding or an open wound is detected, promptly anesthetize the mouse, assess for internal damage, suture the muscle layer again if necessary, and close the skin using surgical clips to facilitate proper wound healing.

Problem 6

Incorrect estradiol injection site (related to step 9).

Potential solution

To address the issue of an incorrect injection site, it is crucial to ensure a proper injection technique. Be vigilant and careful to avoid injecting into surrounding tissues, which could lead to unintended complications.

Resource availability

Lead contact

For further information and requests regarding resources and reagents, please direct your inquiries to the lead contact, Olga Barca-Mayo (olga.barca.mayo@usc.es), and/or the technical contact, María Luengo-Mateos (maria.luengo@rai.usc.es).

Technical contact

For detailed information and support regarding the technical aspects of the research methodology and associated resources, please contact María Luengo-Mateos (maria.luengo@rai.usc.es).

Materials availability

This study did not generate new unique reagents.

Data and code availability

This paper does not report original code.