Development of a Uterosacral Ligament Suspension Rat Model

Beverly J. Miller; Brian K. Jones; Jonathan S. Turner; Steven R. Caliari; Monique H. Vaughan

doi:10.3791/64311

JoVE Journal > Medicine

Medicina

Development of a Uterosacral Ligament Suspension Rat Model

Published: August 17, 2022

doi:

10.3791/64311

Beverly J. Miller¹, Brian K. Jones², Jonathan S. Turner³, Steven R. Caliari^1,3, Monique H. Vaughan⁴

¹Department of Chemical Engineering,University of Virginia, ²Electrophysiology Division,Abbott, ³Department of Biomedical Engineering,University of Virginia, ⁴Department of Obstetrics and Gynecology, Division of Pelvic Medicine and Reconstructive Surgery,University of Virginia

Summary

Pelvic organ prolapse affects millions of women worldwide and yet some common surgical interventions have failure rates as high as 40%. The lack of standard animal models to investigate this condition impedes progress. We propose the following protocol as a model for uterosacral ligament suspension and in vivo tensile testing.

Abstract

Pelvic organ prolapse (POP) is a common pelvic floor disorder (PFD) with the potential to significantly impact a woman’s quality of life. Approximately 10%-20% of women undergo pelvic floor repair surgery to treat prolapse in the United States. PFD cases result in an overall $26.3 billion annual cost in the United States alone. This multifactorial condition has a negative impact on the quality of life and yet the treatment options have only dwindled in the recent past. One common surgical option is uterosacral ligament suspension (USLS), which is typically performed by affixing the vaginal vault to the uterosacral ligament in the pelvis. This repair has a lower incidence of complications compared to those with mesh augmentation, but is notable for a relatively high failure rate of up to 40%. Considering the lack of standard animal models to study pelvic floor dysfunction, there is an urgent clinical need for innovation in this field with a focus on developing cost-effective and accessible animal models. In this manuscript, we describe a rat model of USLS involving a complete hysterectomy followed by fixation of the remaining vaginal vault to the uterosacral ligament. The goal of this model is to mimic the procedure performed on women to be able to use the model to then investigate reparative strategies that improve the mechanical integrity of the ligament attachment. Importantly, we also describe the development of an in situ tensile testing procedure to characterize interface integrity at chosen time points following surgical intervention. Overall, this model will be a useful tool for future studies that investigate treatment options for POP repair via USLS.

Introduction

Pelvic organ prolapse (POP) is a common pelvic floor disorder affecting millions of women worldwide with the potential to significantly impact many aspects of a woman's life, particularly with age¹. Notably, approximately 13% of women in the United States will undergo surgery for prolapse or urinary incontinence². A condition most common after pregnancy and childbirth, prolapse is characterized by the descent of pelvic organs, predominantly the various compartments of the vagina and/or uterus, beyond their normal position in the peritoneal cavity. This leads to bothersome symptoms of vaginal bulge or pressure, bowel, bladder, and sexual dysfunction, and overall reduced quality of life. Other risk factors for POP include obesity, tobacco use, chronic cough, and constipation³.

In healthy women, the pelvic floor organs are supported by the levator ani muscles, uterosacral ligaments (USLs), cardinal ligaments, connective tissue attachments to the pelvic sidewall and the distal structures of the perineal body⁴^,⁵. The USLs are among the most important apical supportive structures for both the uterus and apical vagina, and thus, are often used in surgical correction of POP (Figure 1). Structural support from the USL stems from the dense collagenous connective tissue in the sacral region that transitions into closely packed smooth muscle. Due to this compositional gradient, the USL becomes interwoven with the uterine and vaginal musculature to provide sturdy support for the pelvic organs⁶^,⁷. In the uterosacral ligament suspension (USLS), the USLs are secured to the vaginal vault following a hysterectomy, restoring the vagina and the surrounding structures to their anatomical position in the abdominal compartment. However, regardless of a transvaginal or laparoscopic route, the USLS procedure is plagued by a relatively high failure rate of up to 40% in some studies⁸^,⁹. The recurrence rate of bothersome vaginal bulge symptoms at 5 years post-repair for apical compartment prolapse, such as USLs, was approximately 40% in a large multicenter randomized controlled trial⁹. In the same trial, retreatment for recurrent prolapse at 5 years was approximately 10%. The mechanism of this high failure rate has not been studied, but restoring the vagina and the surrounding structures to their anatomical position requires suture placement in the dense collagenous region of the USL¹⁰^,¹¹ rather than the smooth muscle region. Therefore, the high failure rate could be due to the mechanical and compositional mismatch of the surgically formed vagina-USL interface compared to the complete integration seen in the native cervical-USL attachment.

The economic impact of treating these disorders is also notable, with approximately $300 million spent annually in the US on ambulatory care¹², and more than $1 billion spent annually in direct costs for surgical procedures¹³. Despite the vast economic resources dedicated to treating these conditions, the complications arising from many prolapse surgeries remain discouraging. For example, polypropylene mesh-based apical prolapse repairs, such as sacrocolpopexy, offer higher success rates compared to native tissue repairs¹⁴, but at the cost of potential complications such as mesh exposure or erosion. The FDA received nearly 3,000 complaints related to mesh complications between 2008 and 2010 alone. This culminated in an order by the FDA to halt the manufacture and sale of all transvaginally-placed mesh products for POP in April 2019¹⁵. Therefore, there is a strong clinical need for materials other than polypropylene, and models with which to test them, that may augment native tissue prolapse repairs and increase success rates compared to traditional techniques with suture alone.

Since the FDA announcement in 2019, most pelvic surgeons have stopped using transvaginally-placed mesh for prolapse repairs, prompting investigators to seek new tissue engineering approaches to augment native tissue repairs¹⁶^,¹⁷^,¹⁸ such as with mesenchymal stromal cells (MSCs)⁹^,²⁰. With this shift in focus, there is an urgent need for the refinement of animal models that can assist with development of new materials; the challenge in this process is balancing clinical relevance with cost. To this end, basic science and clinical investigators studying pelvic organ prolapse have taken advantage of several animal models thus far, including rats, mice, rabbits, sheep, swine, and non-human primates¹⁹. The process of identifying an optimal animal model is challenging, as humans are bipedal, have no tail, and have a traumatic birth process compared to other mammalian species²⁰. Swine²¹ have been utilized to simulate robotic sacrocolpopexy, while sheep have been used to simulate vaginal prolapse repairs²². These animal models, while clinically relevant, are limited in feasibility by cost and maintenance. Non-human primates have been used to study the pathogenesis of prolapse; squirrel monkeys in particular are one of the only species other than humans that can develop spontaneous prolapse, making them one of the most relevant animal models²⁰. Non-human primates have also been used to study gynecologic surgical procedures such as sacrocolpopexy²³ and uterine transplantation²⁴. Similar to their sheep and swine counterparts, the primary limitation of non-human primates as an animal model of prolapse is the cost of maintenance, care, and boarding¹⁹.

Although the rodent pelvis is oriented horizontally with a much smaller head-to-birth canal size ratio compared to humans¹⁹, rats are suitable for small animal studies of USLS surgery since they have similar USL anatomy, cellularity, histological architecture, and matrix composition compared to the human USL²⁵. Moreover, they are beneficial in terms of maintenance and boarding. Despite these beneficial attributes, there are no published reports of a rat model of USLS repair. Therefore, the aim is to describe a protocol for hysterectomy and USLS in the multiparous Lewis rat. This protocol will be beneficial for investigators who aim to study the pathophysiology and surgical components of POP using this accessible animal model.

Figure 1: Pelvic organ prolapse. (A) The normal orientation of organs in the peritoneal cavity and (B) the dramatic organ descension when prolapse occurs. Following hysterectomy, (C) uterosacral ligament suspension restores the vagina and surrounding structures to their proper anatomical position. Please click here to view a larger version of this figure.

Protocol

Follow all Institutional Animal Care and Use Committee (IACUC) guidelines, obtaining approval for all animal procedures before beginning. Requirements for aseptic surgery technique can be found from The Guide²⁶ and the Animal Welfare Regulations²⁷. The study was approved by the University of Virginia Institutional Animal Care and Use Committee protocol number 4332-11-20. Obtain multiparous (two litter) female breeders. Rats should be pair housed in a vivarium accredited by the American Association for the Accreditation of Laboratory Animal Care and provided with food and water ad libitum. Animals in this study were Lewis rats obtained from Charles River and were between 4 and 6 months of age to accommodate the two-litter requirement. Animals were maintained on a 12 h light-dark cycle.

1. Pelvic organ prolapse repair using uterosacral ligament suspension

Equipment and surgical area preparation for live animal surgery
1. Prepare the surgical area such that the surgical board is heated to 37 °C using recirculating hot water heating pads along with a sterile waterproof pad. Ensure sterility of the surgical board and surgical area using a bleach free surface disinfectant followed by 70% ethanol wipe.
2. Use autoclave heat sterilization to sterilize all autoclave safe supplies, including surgical instruments, surgical sponges (gauze), cotton swabs, and a disposable drape. Obtain sterile packaged surgical gloves.
3. Obtain electric clippers, ophthalmic ointment, ethanol wipes, cotton swabs, and iodine solution, along with sterile packaged scalpel blade and sutures, and place at the work bench.
Animal preparation for live animal surgery
1. Carefully place the animal into an anesthesia chamber supplied with 2% isoflurane and weigh the animal after the proper plane of anesthesia is reached. Proper anesthetization is confirmed when the animal is non-responsive to a toe pinch.
2. Place the animal onto the surgery board in the prone position with the nose securely in the anesthesia cone supplied with 2% isoflurane. Apply ophthalmic ointment to each of the animals' eyes.
3. Administer opioid analgesic and NSAID analgesic subcutaneously (Table of Materials).
4. Place the animal in the supine position, as shown in Figure 2, and shave off the abdominal fur from the xiphoid process down to the urethral orifice (8 cm x 4 cm). Sterilize the abdomen with three charges of iodine and alcohol to prepare the incision site.
  NOTE: If shaving results in bleeding, achieve hemostasis with pressure prior to preparing the skin with iodine and alcohol prep pad. Maintain iodine on the skin for 30 s.
5. If there is no surgical assistant available, deposit sterile supplies and instruments onto a sterile instrument tray, including sterile cotton swabs, drape(s), sponges (gauze), surgical blade, sutures, and surgical marker (optional). If a surgical assistant is available, then this step can be omitted, and the assistant can provide the sterile instruments following step 1.3.1.
Hysterectomy and uterosacral ligament suspension (USLS)
1. Don a surgical gown, head cover, mask, and sterile gloves. Drape the animal with a sterile field, leaving only the abdomen exposed.
2. Make a 7 cm incision down the linea alba from just below the xiphoid process to the lower nipple line using a scalpel blade. The incision should end ~0.5-1.0 cm rostral from the urethral orifice. Then, make an incision through the muscle layer underneath. Avoid the abdominal wall blood vessel to prevent bleeding.
3. Assemble the abdominal retractor and inspect the abdominal cavity (Figure 3A). Using iris forceps, gently locate the left uterine horn. The uterus is deep insidethe bowel, which is often the structure first encountered upon entering the peritoneal cavity. It is beneficial to first identify the ovary (Figure 3B) and the associated ovarian fat pad.
4. Gently elevate the left uterine horn with a grasper or mosquito clamp and begin hysterectomy by ligating the horn below the ovary and oviduct using a mosquito clamp. The ovaries are delicate structures and are easily damaged or devascularized with manipulation. Take care when elevating the uterine horns; grasp the horn a safe distance from the ovary to achieve this.
5. Continue the hysterectomy by clamping and trimming adjacent vasculature, connective tissue, and fat from the uterine horn using micro scissors. Clamp the connective tissue prior to removal to reduce bleeding. Place the clamps as close to the uterine interface as possible, all the way down to the uterocervical junction (also termed as horn bifurcation).
6. Clamp across the uterine horn near the point of bifurcation using mosquito forceps (Figure 4A-C). Excise the ipsilateral horn just cephalad to the clamp to avoid bleeding. This is located between the utero-cervical junction (just rostral to the cervix) and the utero-tubal ligation point. The vaginal vault will remain post the hysterectomy (Figure 4D).
  NOTE: Due to the small caliber of the rat vessels, ligation of the uterine stumps with a temporary clamp was sufficient for this surgery. However, this technique can be modified as needed with either sealing of the pedicles with electrocautery or suture ligation.
7. Repeat steps 1.3.3-1.3.6 on the right uterine horn to perform a total hysterectomy.
8. Adjust the abdominal retractor to expose the lower pelvis. Inspect the exposed vaginal vault and the pelvic floor support ligamental and connective tissues, which can be seen attached to the vagina and cervix. If possible, identify the ureter bilaterally, which is just medial to the ovaries.
9. Identify the uterosacral ligaments²⁸^,²⁹, shown in Figure 5A, which can be found attached to the cervix just below the remaining stumps of the uterine horns (vaginal vault). The ligament is traced in a cephalad-medial orientation toward the sacrum.
10. Using a 3-0 polydiaxanone suture on a small, tapered needle, place a stitch through the left uterosacral ligament. Place the stitch high on the ligament, close to the sacrum.
11. Tug on the stitch to ensure it has captured the uterosacral ligament-the USL structure inserts into the cervix with the origin diving behind the rectum where it attaches to the sacrum. Again, identify the ureter to ensure it has not been incorporated into or kinked with the uterosacral stitch.
12. Then, pass the left polydiaxanone stitch through the left aspect of the vaginal vault (Figure 5B), with care to incorporate both the anterior and posterior aspects of the vaginal cuff. Repeat the steps to complete the USLS procedure on the right side. Multiple stitches can be placed bilaterally, if desired.
13. Once the uterosacral stitches are placed bilaterally, securely tie the suture using a square knot, as shown in Figure 5C, such that the vaginal vault is elevated cephalad toward the sacrum; this completes the uterosacral ligament suspension.
Closing the surgical wound
1. Replace the abdominal contents back into their anatomical position within the peritoneal cavity. Close the deep layers of the abdominal wall (peritoneum, fascia, muscle) with continuous suture pattern of 4-0 to 6-0 polyglactin 910 or polydiaxanone suture.
2. Close the skin with a running subcuticular (or interrupted) stitch of 4-0 to 6-0 polydiaxanone or polyglactin 910. Administer antibiotic subcutaneously as needed for surgical site infection prophylaxis.
3. Perform post-surgical monitoring until the animal has regained sufficient consciousness to maintain sternal recumbency. Do not return the animal to social housing until fully recovered.

Figure 2: Animal preparation for live surgery. Removing fur from the area surrounding the incision site is necessary for proper aseptic technique. The area shown in panels (A) and (B) are guidelines. Researchers should remove enough hair such that sterile instruments make no contact with hair during surgery. Please click here to view a larger version of this figure.

Figure 3: Preserving the ovaries. The uterine horns are typically not visible when the abdomen is first opened, as shown in (A). Once a horn is located and followed to find (B) the ovary and oviduct where they connect to the horn, the top of the horn can be clamped, and the horn separated to begin hysterectomy. Please click here to view a larger version of this figure.

Figure 4: Removing the uterine horns. Hysterectomy in the rat involves (A) both uterine horns (B) clamped at the uterocervical junction and (C) excised. The vaginal vault from each horn remains with the (D) cervical/uterine stump (arrow) connecting them. Please click here to view a larger version of this figure.

Figure 5: Uterosacral ligament suspension. (A) Orientation of the uterosacral ligaments in relation to the created vaginal vault structures. When placing sutures for the uterosacral ligament suspension (USLS) repair, (B) sutures capture the uterosacral ligament and then pass through both the anterior and posterior aspects of the vaginal cuff. (C) Secured to the uterosacral ligament, the vaginal vault is now elevated cephalad toward the sacrum. Please click here to view a larger version of this figure.

2. Uniaxial tensile testing

NOTE: The testing system and software used was operated following the manufacturer's guidelines for calibration and testing. All testing occurred at 22 °C.

Specimen preparation
1. Euthanize the rat using an IACUC-approved pharmacological procedure. Ensure death via secondary physical method. Here, CO₂ inhalation was used followed by cardiac puncture. Expose the vaginal vault in preparation for tensile mechanical testing. In the current study, perform tensile testing on native uterosacral ligaments (control), as well as on animals who had undergone uterosacral ligament suspension as described above (POP).
2. Test ligaments in situ 24 weeks following surgery. A terminal timepoint of minimum 8 weeks is suggested to allow for the complete reabsorption of the sutures.
  1. Following humane euthanasia, make an incision down the linea alba to expose the abdomen.
  2. Begin dissecting off the adipose tissue until the vaginal vault is visible. Continue to dissect off the abdominal fat pads until the intact USLs is clearly visible (control animals, Figure 6A) or the junction between the uterosacral ligament and the vaginal vault is visible (POP animals, Figure 6C). Use caution to not pull on the junction to remove adipose tissue, but rather using careful cuts with micro-scissors to maintain consistency between samples.
  3. Using a flexible ruler, measure the distance between the uterosacral insertion (posterior to the rectum) and the vaginal vault. This value is the original length of the tissue.
    NOTE: The original length of the tissue, the gauge length, for control USLs measured 13.4 ± 0.5 mm while the gauge length for USLs repair measured 12.8 ± 0.4 mm.
  4. Thread umbilical tape behind the intact USL (control, Figure 6B) or the USLS junction (POP, Figure 6D) such that the tissue is centered on the umbilical tape. Measure the height and width of the tissue where it intersects with the umbilical tape using digital calipers. These values will be used to calculate cross-sectional area.
  5. Attach a large compression plate (Table of Materials) via the base adapter and position the animal atop such that the specimen is centered beneath the grip holder.
Tensile testing
1. Program the tensile testing regime into the software: pre-load, pre-condition, pull to failure. This follows previous pelvic floor²⁹ and reproductive tissue³⁰ mechanical testing protocols.
2. Set up the instrument in preparation of tensile testing. For the current study, use a 10 N load cell, a 3D printed grip, and a base adapter to attach a compression platen as shown in Figure 7.
  NOTE: Any base set-up that can support the full size of the animal is acceptable. Use any grip that can securely hold the umbilical tape. A custom 3D printed holder and grip from previous studies³¹^,³² was used in this testing. STL files were included as supplemental files.
  1. Position the animal such that the specimen is centered beneath the grip (Figure 8A). Immobilize the pelvic region surrounding the specimen by securing the animal to the platen (Figure 8B).
  2. Lower the load cell such that the tails of the umbilical tape easily reach the grip. Secure the umbilical tape in the grip, leaving the tape slack to avoid specimen manipulation.
3. Open the pre-conditioning test in the software interface and label the test with the sample name. Ensure that the pre-conditioning method includes the pre-load step.
4. Click to start the pre-conditioning test, which will pre-load the sample at 0.015 N. Once the pre-load force is stable, the test will precondition the sample at an elongation rate of 0.1 mm/s for 30 s. Allow the tissue to rest for 1 min. While waiting, load the pull-to-failure testing regime.
  NOTE: The pre-load force may vary depending on the instrument limitations and the testing conditions. Refer to previous studies where the reported pre-load ranges from 0.015 N to 0.1 N²⁹^,³³^,³⁴^,³⁵^,³⁶.
5. Open the testing regime that is programmed to pull to failure. Label the test with the sample name and click on Okay to get to the next window. Input the gauge length of the sample and then click on Next to transition to the test page.
6. Balance all and click on Start. Allow the test to run at an elongation rate of 0.1 mm/s until the tissue has been pulled to failure. The test will produce load-displacement data.
Calculation of stress, strain, and modulus for tensile testing
1. Using the load-displacement data, the cross-sectional area, and the gauge length from the sample, calculate the stress (MPa) and strain (%) as previously reported³⁷^,³⁸^,³⁹^,⁴⁰^,⁴¹. Use Equation 1 and Equation 2 shown below. Note that stretching of the tape during testing should also be accounted for in these calculations.
  Equation 1
  Equation 2
  1. From the load-displacement curve (Figure 9A,D), calculate the stiffness (linear slope, N/mm) and ultimate load. From the stress strain curve, calculate the tangent modulus (linear slope, MPa) and the ultimate stress. The linear region of the stress strain curve is noted in Figure 9B,E with the calculated tangent modulus from this region shown in Figure 9C,F for both experimental groups.
    NOTE: For both the stiffness and tangent modulus, identify the linear portion by choosing a window of points that maximizes the R² value for a linear regression³⁷^,⁴¹.

Figure 6: Specimen preparation for uniaxial tensile testing. (A) The exposed control USLs before (B) the umbilical tape is threaded behind the tissue. (C) USL-vaginal vault junction after the complete dissolution of the sutures with (B) the umbilical tape threaded behind the tissue in preparation of tensile testing. Please click here to view a larger version of this figure.

Figure 7: The mechanical testing system. (A) The testing system in tensile testing mode used with (B) 3D printed holder and (C) 3D printed sample grip complete with a textured strip to improve grip. Configuration of the pieces shown in panel (D). Please click here to view a larger version of this figure.

Figure 8: Set-up of the tensile testing. (A) The specimen is centered beneath the grip and holder. (B) The animal and tissue surrounding the specimen are held stationary prior to the start of the tensile test. As shown by the inset image, securing the surrounding tissue is essential to isolate the tissue of interest. Please click here to view a larger version of this figure.

Figure 9: Example of tensile test data output and analysis. (A) The load-displacement curve for a control sample followed by (B) the stress strain analysis and (C) the slope of the line curve fit equation showing the tangent modulus in MPa. (D–F) shows the same process for a USLS sample. Please click here to view a larger version of this figure.

Representative Results

Surgical feasibility and uterosacral suture placement
There were no intraoperative complications related to hysterectomy or uterosacral ligament suspension in any of the animals. There was minimal bleeding during removal of the uterine horns, provided the adjacent vasculature was clamped prior to removal. Limited bleeding allowed for good visualization of the uterosacral ligaments for suture placement and prevented intra-operative bowel, rectum, ureteral, or bladder injury. Following placement of the sutures, the newly formed USL-vaginal vault junction prevented movement of the cervical/uterine stump as shown in Figure 5C. During the first three postoperative days, the animals were checked on daily, and then bi-weekly basis until the end of the experiment. With the extended-release opioid and NSAID analgesics administered at the time of surgery, additional analgesics were found to be unnecessary. Based on our experience with 16 animal surgeries (n = 8 for both control and USLS groups), a drop in weight should be expected in the first week following surgery with an average loss of 5.7 ± 1.4% from surgery day weight. As expected, the rats slowly gained weight over the subsequent 23 weeks, with an average weight gain of 15.1 ± 4.5% over the course of the experiment.

Mechanical testing of the USLS repair
To demonstrate the functionality of the USLS repair, uniaxial tensile testing was performed. After euthanasia of the animal at the chosen post-operative timepoint, 24 weeks in this study, the surgical area should be carefully dissected to visualize the USL-vaginal vault junction as shown in Figure 6A. Compared to other methodologies for testing the rat USLs together with other supportive structures and pelvic organs²⁹^,⁴², the method described here is the first to test the rat USL in an isolated manner. The umbilical tape used in this study was strategically chosen for its flexibility as the tape compliance allowed for minimal disruption of the tissue during tensile testing preparation. Load displacement data, therefore, must be adjusted to account for the small amount of stretch contributed by the umbilical tape. Figure 9 provides an example of data obtained via tensile testing with Figure 9A providing an example of a typical stress-strain plot. Reporting of stress-strain data is recommended as this information is normalized and independent of the size of the specimens³⁴ and can be better compared across studies. For the intact uterosacral ligament, we report structural properties such as ultimate load (2.9 ± 0.5 N) and stiffness (0.4 ± 0.1 N/mm) as well as normalized material properties such as ultimate stress (2.1 ± 0.4 MPa), ultimate strain (1.6 ± 0.5) and tangent modulus (4.0 ± 1.1 MPa). In the uniaxial tests performed on the rat reproductive organs and all their supportive tissue connections by Moalli et al., they reported an ultimate load at failure (13.2 ± 1.1 N) and stiffness (2.9 ± 0.9 N/mm) higher than the isolated USL²⁹. The work done by Moalli et al. and other literature³⁴^,³⁵ mention the high variability between tested specimens as shown in the data presented here. For the uterosacral ligament suspension repair, we found all structural material properties (stiffness, 0.33 ± 0.13 N/mm; ultimate load, 2.6 ± 1.3 N) and normalized material properties (ultimate stress, 1.8 ± 0.7 MPa; ultimate strain 1.3 ± 0.3; tangent modulus, 3.0 ± 0.9 MPa) to be lower than that of the native USL.

Discussion

The protocol is notable for several advantages. To our knowledge, it is the first published description of USLS in the rat model and will provide future investigators with reproducible steps for performing this procedure in the research setting. Second, we include a novel protocol for tensile testing of the native and surgical interface of the USL. The tensile testing protocol could be utilized in similar studies that investigate new tissue engineering approaches to augment native tissue repairs such as USLS. Moreover, the rat model itself is useful for the study of pelvic floor disorders due to ease of handling/boarding, short lifespan, and cost efficiency compared to larger animal models. Limitations of the protocol include an inability to assess one of the main complications of USLS, ureteral kinking. Despite this, we had no cases of presumed ureteral injury in this study. Another consideration is that the horizontal orientation of the pelvis, small fetal head-to-birth canal ratio, and lack of spontaneous prolapse in the rat model does limit some applicability of results to humans. However, the use of multiparous rats is a strength of this study since this accounts for the leading risk factor in the development of POP³.

The establishment of a successful protocol for hysterectomy and USLS in the Lewis rat will be a useful tool for future researchers investigating surgical components of POP, while minimizing variability in testing the mechanical behavior of the USL. Surgical animal models are beneficial in that they allow researchers to design clinically relevant experiments that control for parity, body mass, disease, and nutrition³⁴ while mitigating the ethical risk of initial study in humans. Further, standardized models for POP allow researchers to bypass the limitations of human tissue collection. In particular, the tensile testing methods described in this protocol will enable consistency between studies. Previous rodent models tested the mechanical properties of the entire pelvic region, which includes the cervix, vagina, and the multiple pelvic support ligaments²⁹^,⁴². The methods described here allow for measurement of the USL in a way that maintains the native spinal and cervical attachments. It should be noted that the tensile testing methods do not assess the USL alone, but rather the USL in combination with its insertion at the sacrum and cervix. This is a strength of the study as it reflects the usual in situ forces to which the ligament is subjected. We acknowledge that the mechanical behavior of the isolated ligament would be different if it were tested ex vivo without its native attachments. This is especially true since the rat structures are small and limit the feasibility of collecting a sample suitable for ex vivo testing. The USLs do experience loading in multiple directions in situ, so the uniaxial nature of the test is a limitation, but using this method allows for meaningful comparisons between previous studies of rat USL mechanics²⁹^,⁴². While there is currently no widely accepted standard mechanical testing protocol, this model will be a useful tool for future tissue engineering studies in the field.

Several steps described in this protocol are critical to the health and well-being of the animals as well as the reproducibility of the USLS surgery and subsequent tensile testing. First, it is essential to obtain both the analgesic and the anti-inflammatory drugs described as the analgesic alone was found to be inadequate for pain management. The prophylactic antibiotic decreases the risk of surgical site infection and is the standard of care in human surgery. Regarding the USLS surgical procedure, avoiding damage to the ovaries and minimizing blood loss are essential for a successful surgery. Steps 1.3.3 and 1.3.4 describe separating the top of the uterine horn from the adjacent ovary; care should be taken to maintain this dissection on the side of the uterine horn to prevent disruption of delicate vessels around the ovary, which can result in excessive bleeding. Of note, other investigators have shown that ovarian function is preserved after removal of the uterine horns⁴³. Moreover, if the ovaries are disrupted or removed, the overall collagen fibril architecture will be disturbed, altering the mechanical properties of its tissues⁴⁴^,⁴⁵. Once the uterine horn is safely separated from the ovary, there is a clear plane of dissection allowing isolation of the uterine horn from the surrounding fat pads and vasculature. Despite the clear plane of dissection, the pedicles along the uterine horn should be secured with a clamp prior to transection with micro scissors. Contrary to surgical practice in humans, we have found that suture ligation of the hysterectomy pedicles is unnecessary, as clamping the pedicle prior to transection ensures adequate hemostasis. Step 1.3.6 of the protocol describes this careful process to minimize blood loss. As the hysterectomy is being performed, great care should be taken to identify the ureters as mentioned in steps 1.3.6 and 1.3.8. Understanding the anatomical proximity of the ureter is critical, as one of the most common complications associated with the USLs in humans is ureteral injury⁴⁶.

In conclusion, we present a novel protocol for performing hysterectomy, uterosacral ligament suspension, and tensile testing of the USL in a rat model. We anticipate that our findings will assist future basic science investigators by providing a clear, reproducible description of these procedures and thereby allow for advancement of pelvic organ prolapse research.

Declarações

The authors have nothing to disclose.

Acknowledgements

We thank Prof. Silvia Blemker for use of her Instron and Prof. George Christ for use of his surgical space as well as the 3D printed holder and grip. This work was supported by the UVA-Coulter Translational Research Partnership and the DoD (W81XWH-19-1-0157).

Materials

Alcohol prep pad	BD	326895
Artificial Tear Ointment	American Health Service Sales Corp	PH-PARALUBE-O
Bluehill software	Instron	Bluehill 3
Cavicide 1 disinfectant	Fisher Scientific	22 998 800
Compression platean	Instron	2501-163
Cotton swabs	Puritan Medical	806-WC
Gauze Sponge, 8-Ply	VWR	95038-728
Mosquito Forceps	Medline Industries	MMDS1222115
Needle Holder	Medline Industries	DYND04045
Operating Scissors, 5½", Sharp	American Health Service Sales Corp	4-222
Opioid Analgesic (Buprenorphine XR)	Fidelis Animal Health	Ethiqa XR	0.65 mg/kg SC Q72
NSAID Analgesic (Meloxicam SR)	Wildlife Pharmaceuticals, LLC	Meloxicam SR	1 mg/kg SC q72
PDS II, 3-0 Polydioxanone Suture, SH-1	Ethicon	Z316H
PDS II, 5-0 P olydioxanone Suture, RB-1	Ethicon	Z303H
Retractor	Medline Industries	MDS1862107
Scalpel Blade Stainless Surgical #10	Miltex	4-310
Scalpel Handle	Medline Industries	MDS15210
Scissor, Micro, Curved, 4.5"	Westcott	MDS0910311
Single Column Universal Testing System	Instron	5943 S3873	1 kN force capacity, 10 N load cell
Sterile Natural Rubber Latex Gloves	Accutech	91225075
Suture,Vicryl,6-0,P-3	Ethicon	J492G
Tape,Umbilical,Cotton,1/8X18"	Ethicon	U10T
Tension and Compression Load Cell	Instron	2530-10N	10N load cell (1 kgf, 2 lbf)
Veterinary surgical adhesive (skin glue)	Covetrus	31477

Referências

Olsen, A. L., et al. Epidemiology of surgically managed pelvic organ prolapse and urinary incontinence. Obstetrics and Gynecology. 89 (4), 501-506 (1997).
Wu, J. M., et al. Lifetime risk of stress urinary incontinence or pelvic organ prolapse surgery. Obstetrics and Gynecology. 123 (6), 1201-1206 (2014).
Kenton, K., Mueller, E. R. The global burden of female pelvic floor disorders. BJU International. 98, 1-7 (2006).
Herschorn, S. Female pelvic floor anatomy The pelvic floor, supporting structures, and pelvic organs. Reviews in Urology. 6, 2-10 (2004).
Jelovsek, J. E., Maher, C., Barber, M. D. Pelvic organ prolapse. The Lancet. 369 (9566), 1027-1038 (2007).
Campbell, R. M. The anatomy and histology of the sacrouterine ligaments. American Journal of Obstetrics and Gynecology. 59 (1), 1-12 (1950).
Reisenauer, C., et al. The role of smooth muscle in the pathogenesis of pelvic organ prolapse – An immunohistochemical and morphometric analysis of the cervical third of the uterosacral ligament. International Urogynecology Journal and Pelvic Floor Dysfunction. 19 (3), 383-389 (2008).
Lavelle, R. S., Christie, A. L., Alhalabi, F., Zimmern, P. E. Risk of prolapse recurrence after native tissue anterior vaginal suspension procedure with intermediate to long-term followup. Journal of Urology. 195 (4), 1014-1020 (2016).
Jelovsek, J. E., et al. Effect of uterosacral ligament suspension vs sacrospinous ligament fixation with or without perioperative behavioral therapy for pelvic organ vaginal prolapse on surgical outcomes and prolapse symptoms at 5 years in the OPTIMAL randomized clinical trial. JAMA – Journal of the American Medical Association. 319 (15), 1554-1565 (2018).
Bradley, M. S., et al. Vaginal uterosacral ligament suspension: A retrospective cohort of absorbable and permanent suture groups. Female Pelvic Medicine & Reconstructive Surgery. 24 (3), 207-212 (2018).
Cola, A., et al. Native-tissue prolapse repair: Efficacy and adverse effects of uterosacral ligaments suspension at 10-year follow up. International Journal of Gynecology and Obstetrics. , (2022).
Sung, V. W., Washington, B., Raker, C. A. Costs of ambulatory care related to female pelvic floor disorders in the United States. American Journal of Obstetrics and Gynecology. 202 (5), 1-4 (2010).
Subak, L. L., et al. Cost of pelvic organ prolapse surgery in the United States. Obstetrics and Gynecology. 98 (4), 646-651 (2001).
Siddiqui, N. Y., et al. Mesh sacrocolpopexy compared with native tissue vaginal repair: A systematic review and meta-analysis. Obstetrics & Gynecology. 125 (1), 44-55 (2015).
FDA takes action to protect women’s health, orders manufacturers of surgical mesh intended for transvaginal repair of pelvic organ prolapse to stop selling all devices. FDA News Release Available from: https://www.fda.gov/news-events/press-announcements/fda-takes-action-protect-womens-health-orders-manufacturers-surgical-mesh-intended-transvaginal (2019)
Brincat, C. A. Pelvic organ prolapse reconsidering treatment, innovation, and failure. JAMA – Journal of the American Medical Association. 322 (11), 1047-1048 (2019).
Cundiff, G. W. Surgical innovation and the US Food and Drug Administration. Female Pelvic Medicine & Reconstructive Surgery. 25 (4), 263-264 (2019).
Luchristt, D., Weidner, A. C., Siddiqui, N. Y. Urinary basement membrane graft-augmented sacrospinous ligament suspension: a description of technique and short-term outcomes. International Urogynecology Journal. 33 (5), 1347-1350 (2022).
Couri, B. M., et al. Animal models of female pelvic organ prolapse: Lessons learned. Expert Review of Obstetrics and Gynecology. 7 (3), 249-260 (2012).
Mori da Cunha, M. G. M. C., et al. Animal models for pelvic organ prolapse: systematic review. International Urogynecology Journal. 32 (6), 1331-1344 (2021).
Kasabwala, K., Goueli, R., Culligan, P. J. A live porcine model for robotic sacrocolpopexy training. International Urogynecology Journal. 30 (8), 1371-1375 (2019).
Mansoor, A., et al. Development of an ovine model for training in vaginal surgery for pelvic organ prolapse. International Urogynecology Journal. 28 (10), 1595-1597 (2017).
Liang, R., et al. Impact of prolapse meshes on the metabolism of vaginal extracellular matrix in rhesus macaque. American Journal of Obstetrics and Gynecology. 212 (2), 1-7 (2015).
Johannesson, L., et al. Preclinical report on allogeneic uterus transplantation in non-human primates. Human Reproduction. 28 (1), 189-198 (2013).
Iwanaga, R., et al. Comparative histology of mouse, rat, and human pelvic ligaments. International Urogynecology Journal. 27 (11), 1697-1704 (2016).
National Research Council. . Guide for the Care and Use of Laboratory Animals: Eighth Edition. , (2011).
Federal Animal Welfare Regulations. National Archives Available from: https://www.ecfr.gov/current/title-9/chapter-l/subchapter-A/part-2/subpart-C/section-2.31 (2022)
Ma, Y., et al. Knockdown of Hoxa11 in vivo in the uterosacral ligament and uterus of mice results in altered collagen and matrix metalloproteinase activity. Biology of Reproduction. 86 (4), 100 (2012).
Moalli, P. A., et al. A rat model to study the structural properties of the vagina and its supportive tissues. American Journal of Obstetrics and Gynecology. 192 (1), 80-88 (2005).
Yoshida, K., et al. Mechanics of cervical remodelling: Insights from rodent models of pregnancy. Interface Focus. 9 (5), 20190026 (2019).
Christ, G. J., Sharma, P., Hess, W., Bour, R. . Modular biofabrication platform for diverse tissue engineering applications and related method thereof. , (2020).
Smith, K., Christ, G. J. . Incorporation of in vitro double seeding for enhanced development of tissue engineered skeletal muscle implants. , (2019).
Becker, W. R., De Vita, R. Biaxial mechanical properties of swine uterosacral and cardinal ligaments. Biomechanics and Modeling in Mechanobiology. 14 (3), 549-560 (2015).
Donaldson, K., Huntington, A., De Vita, R. Mechanics of uterosacral ligaments: Current knowledge, existing gaps, and future directions. Annals of Biomedical Engineering. 49 (8), 1788-1804 (2021).
Baah-Dwomoh, A., McGuire, J., Tan, T., De Vita, R. Mechanical properties of female reproductive organs and supporting connective tissues: A review of the current state of knowledge. Applied Mechanics Reviews. 68 (6), 1-12 (2016).
Tan, T., Cholewa, N. M., Case, S. W., De Vita, R. Micro-structural and biaxial creep properties of the swine uterosacral-cardinal ligament complex. Annals of Biomedical Engineering. 44 (11), 3225-3237 (2016).
Kurtaliaj, I., Golman, M., Abraham, A. C., Thomopoulos, S. Biomechanical testing of murine tendons. Journal of Visualized Experiments. (152), e60280 (2019).
Griffin, M., et al. Biomechanical characterization of human soft tissues using indentation and tensile testing. Journal of Visualized Experiments. (118), e54872 (2016).
Feola, A., et al. Parity negatively impacts vaginal mechanical properties and collagen structure in rhesus macaques. American Journal of Obstetrics and Gynecology. 203 (6), 1-8 (2010).
Tan, T., et al. Histo-mechanical properties of the swine cardinal and uterosacral ligaments. Journal of the Mechanical Behavior of Biomedical Materials. 42, 129-137 (2015).
Abramowitch, S. D., Feola, A., Jallah, Z., Moalli, P. A. Tissue mechanics, animal models, and pelvic organ prolapse: A review. European Journal of Obstetrics and Gynecology and Reproductive Biology. 144, 146-158 (2009).
Lowder, J. L., et al. Adaptations of the rat vagina in pregnancy to accommodate delivery. Obstetrics and Gynecology. 109 (1), 128-135 (2007).
Koebele, S. V., et al. Hysterectomy uniquely impacts spatial memory in a rat model: A role for the nonpregnant uterus in cognitive processes. Endocrinology. 160 (1), 1-19 (2019).
Kafantari, H., et al. Structural alterations in rat skin and bone collagen fibrils induced by ovariectomy. Bone. 26 (4), 349-353 (2000).
Daghma, D. E. S., et al. Computational segmentation of collagen fibers in bone matrix indicates bone quality in ovariectomized rat spine. Journal of Bone and Mineral Metabolism. 36 (3), 297-306 (2018).
Manodoro, S., Frigerio, M., Milani, R., Spelzini, F. Tips and tricks for uterosacral ligament suspension: how to avoid ureteral injury. International Urogynecology Journal. 29 (1), 161-163 (2018).

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Miller, B. J., Jones, B. K., Turner, J. S., Caliari, S. R., Vaughan, M. H. Development of a Uterosacral Ligament Suspension Rat Model. J. Vis. Exp. (186), e64311, doi:10.3791/64311 (2022).