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Isolation and Characterization of the Murine Uterosacral Ligaments and Pelvic Floor Organs

Published: March 3, 2023 doi: 10.3791/65074


This article presents a detailed protocol for dissecting uterosacral ligaments and other pelvic floor tissues, including the cervix, rectum, and bladder in mice, to expand the study of female reproductive tissues.


Pelvic organ prolapse (POP) is a condition that affects the integrity, structure, and mechanical support of the pelvic floor. The organs in the pelvic floor are supported by different anatomical structures, including muscles, ligaments, and pelvic fascia. The uterosacral ligament (USL) is a critical load-bearing structure, and injury to the USL results in a higher risk of developing POP. The present protocol describes the dissection of murine USLs and the pelvic floor organs alongside the acquisition of unique data on the USL biochemical composition and function using Raman spectroscopy and the evaluation of mechanical behavior. Mice are an invaluable model for preclinical research, but dissecting the murine USL is a difficult and intricate process. This procedure presents an approach to guide the dissection of murine pelvic floor tissues, including the USL, to enable multiple assessments and characterization. This work aims to aid the dissection of pelvic floor tissues by basic scientists and engineers, thus expanding the accessibility of research on the USL and pelvic floor conditions and the preclinical study of women's health using mouse models.


Approximately 50% of women are affected by pelvic organ prolapse (POP)1,2. About 11% of these women fit the criteria for undergoing surgical repair, which has a poor success rate (~30%)3,4. POP is characterized by the descent of any or all of the pelvic organs (i.e., bladder, uterus, cervix, and rectum) from their natural position due to the failure of the USL and the pelvic floor muscles to provide adequate support5. This condition involves anatomical dysfunction and disruption of the connective tissue, as well as neuromuscular injury, in addition to predisposing factors3,6. POP is associated with multiple factors such as age, weight, parity, and delivery type (i.e., vaginal or caesarian births). These factors are thought to affect the mechanical integrity of all the pelvic floor tissues, with pregnancy and parity thought to be the main drivers of POP5,7,8.

The uterosacral ligaments (USLs) are important supportive structures for the uterus, cervix, and vagina and tether the cervix to the sacrum4. Damage to the USLs puts women at increased risk of developing POP. It is believed that pregnancy and childbirth impose additional strain on the USL, which potentially induces injury and increases the chances of POP. The USL is a complex tissue composed of smooth muscle cells, blood vessels, and lymphatics distributed heterogeneously along the ligament, which can be divided into three distinct sections: cervical, intermediate, and sacral region9. The mechanical integrity of the USL is derived from extracellular matrix (ECM) components like collagens, elastin, and proteoglycans5,9,10. Type I collagen fibers are known to be a major load-bearing tensile component of ligamentous tissues and are, therefore, likely involved in USL failure and POP11.

There is a lack of knowledge regarding the causes, prevalence, and effects of POP in women. The development of an appropriate animal model of POP is necessary to advance our understanding of the female pelvic floor. Mice and humans have similar anatomical landmarks within the pelvis, such as the ureters, rectum, bladder, ovaries, and round ligaments9, as well as similar intersection points of the USL with the uterus, cervix, and sacrum. Further, mice offer ease of genetic manipulation and have the potential to be an easily accessible, cost-effective model for the study of POP9.

This study developed a method to access and isolate the USL and the different pelvic floor tissues from nulliparous (i.e., never pregnant) mice. The extracted USLs were subjected to enzymatic digestion (i.e., to remove collagens and glycosaminoglycans), tested to determine the mechanical response under tensile loading, and evaluated for biochemical composition in a proof-of-concept study. The ability to isolate intact tissues will facilitate further mechanical and biochemical characterizations of the pelvic floor components, which is a crucial first step toward improving our understanding of the injury risks related to childbirth, pregnancy, and POP.

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All animal experiments and procedures were performed according to protocol #2705, approved by the Animal Care and Use Committee of the University of Colorado Boulder. Six week old female C57BL/6J mice were used for the present study. The animals were obtained from a commercial source (see Table of Materials).

1. Animal preparation

  1. Euthanize the animal following the institutionally approved method.
    NOTE: The present study used CO2 inhalation in alignment with the American Veterinary Medical Association's guidelines (a displacement rate of 30% to 70% of the chamber volume with CO2 per minute), followed by cervical dislocation, to ensure successful euthanization.
    1. Work under a hood, if possible, to minimize the spread of mice allergens. Once the mouse stops moving and breathing, allow 2 min or more to verify the lack of response.
      NOTE: If the mouse is pregnant or post-partum, the pups must be individually euthanized. Pups E15.5 and older must be decapitated during the dissection.
  2. Prepare the dissection setup with a dissection pad, an 11-blade scalpel, curved thin sharp scissors, two pairs of forceps, curved forceps, 5-0 polyglactin suture, a dissecting microscope, and six pins (Figure 1, see Table of Materials).
  3. Place the mouse on the pad, and pin the forelimbs down (Figure 2A). Make an incision of approximately 1-1.5 cm in the abdomen with scissors (Figure 2B). Gently use the scissors to separate the skin at the cranial, caudal, and lateral sides of the incision (Figure 2C, D).
  4. Flip the mouse to its dorsal side, and gently peel back the skin toward the hindlimbs to remove the skin away from the dissection site (Figure 2E-H).
  5. Pin the mouse at the limbs (Figure 2I), and make an incision of approximately 1 cm into the abdomen from the thorax to the pelvis (Figure 2J).
    NOTE: Ensure not to damage the underlying organs.
  6. Gently push the organs toward the thorax to clear the field of view (Figure 2K).
    NOTE: Irrigate the tissues with 1x PBS to maintain hydration.
  7. Clear all of the fat tissue from the pelvic floor (Figure 2L-N).
    ​NOTE: Use forceps to gently pull and clear fat off the organs and tissues of interest.

Figure 1
Figure 1: A clean workspace with all the tools needed to perform the dissections. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Removal of the skin and opening of the pelvic and thoracic cavities of the mouse. (A) Pinning down all the limbs. (B) Initial incision. (C) Separating the skin from underlying fascia using scissors. (D) Cutting of the skin and preparing for removal. (E-G) Pulling the skin off by going around the mouse. (H) Completely removing the skin from the dorsal side. (I) Complete removal of the skin from the torso, and re-pinning of the mouse limbs. (J) Opening of the abdomen. (K) View of the open abdomen. (L) Moving the organs out of the field of view. (M) Removing the fat. (N) View of the cleared pelvic floor. Please click here to view a larger version of this figure.

2. USL harvesting

  1. Cut the uterine horns from the ovaries (Figure 3B), pull away from the field of view, and cut off at the cervical connection (Figure 3C).
    NOTE: The uterine horns can be identified following the schematic in Figure 3A. Irrigate the tissues with 1x PBS to maintain hydration.
  2. Cut the ureters away from the bladder connection (Figure 3D).
    NOTE: This is to avoid confusion with the USL.
  3. Cut the colon as close to the cervix as possible (Figure 3E,F).
    NOTE: Irrigate the tissues with 1x PBS to maintain hydration.
  4. Place the mouse along with the dissection pad under the dissecting scope to visualize the USLs (Figure 3G).
  5. Gently use the forceps to clean the surrounding fat from the USLs.
    NOTE: Use a second pair of forceps to hold the cervix up at a small angle to enhance the visualization of where the USL intersects with the cervix. Irrigate the tissues with 1x PBS to maintain hydration.
  6. Tie a 5-0 polyglactin suture around the cervical end of both USLs (Figure 4B, C).
    NOTE: The USLs can be identified using the schematic and magnification images (Figure 4I-K)
  7. In this study, one USL is used for morphological or biochemical analyses (i.e., Raman microscopy, immunohistochemistry, histology). Cut the cervical end of the USL, leaving a piece of the cervix attached, and cut a piece of muscle from the bottom of the USL (Figure 4D). Place the dissected tissue in a bath with 1x PBS to keep the tissue hydrated (Figure 4G, H).
  8. Use the remaining USL for mechanical testing and imaging. Cut the cervical end of the USL, leaving a piece of cervix attached, to facilitate the mechanical setup (Figure 4D).
    NOTE: The cervical tissue will act as an anchor to secure the USL during the mechanical test.
  9. Once all the tissues of interest are harvested (steps 3-5), dislocate the femurs from the pelvis (Figure 4E).
    NOTE: One should hear a faint clicking sound when the femoral head is disarticulated from the acetabular cup.
  10. Cut the pelvic bone from the distal and proximal ends of the pelvic bone, leaving about 10 mm of total tissue (Figure 4F). Place the dissected tissue in 1x PBS.

Figure 3
Figure 3: Cleared pelvic floor for USL dissection. (A) Schematic of the anatomy. (B) Cutting the uterine horns at the ovarian connection. (C) Cutting off uterine horns. (D) Cutting of the ureters. (E) Cutting of the colon. (F) A clear view of the rectum and USLs. (G) Placing the mouse and dissection pad under the dissecting scope. Please click here to view a larger version of this figure.

Figure 4
Figure 4: View of the USL and surrounding tissues and dissection of the USLs. (A) Schematic of anatomical landmarks surrounding the USL. (B) Tying a suture around the cervical ends. (C) Slicing off the cervical ends of the USL. (D) Slicing off the USL to be used for the biochemical analyses at the sacral connection. (E) Cutting of the femurs from the pelvic bone. (F) Cutting off the proximal end of the pelvis. (G) Dissecting the USL in a 35 mm Petri dish. (H) The USL with the attached pelvis in a 35 mm petri dish. (I) The USL and rectum at 0.75x magnification. (J) Removing fat from the USL. (K) Cleaning of the USLs at 1.0x magnification. Scale bar = 2 mm. Please click here to view a larger version of this figure.

3. Bladder harvesting

  1. After the fat is cleared, hold the bladder with the forceps, and gently lift it at an angle of approximately 40° (Figure 5A).
  2. With the scissors, slice off the bladder from the distal side, right above the cervix (Figure 5B).
  3. Place the tissue in a bath with 1x PBS to keep the tissue hydrated (Figure 5H).

4. Rectum harvesting

  1. Once the USLs are disconnected from the cervix and the bladder is dissected, lift the cervix at an angle of approximately 40° with the forceps. There is the rectovaginal fascia that connects the rectum and the cervix. With the scalpel, gently cut this connection (Figure 5C, D).
  2. Cut the pubic bone at the pubic symphysis using scissors. Gently widen the workspace to increase visual access to the tissue insertions.
  3. With the forceps, gently pull the rectum toward the thorax, and use the scissors to follow the rectum from its posterior side to the anus. Cut the rectum at the anus (Figure 5E).
  4. Place the tissue in 1x PBS to keep the tissue hydrated (Figure 5I).

5. Cervix-vagina complex harvesting

  1. After the USLs are removed from the cervix, use the forceps to hold the cervix. Cut the cervix as close to the vulva as possible using scissors (Figure 5F, G).
    NOTE: Ensure to cut the pubic symphysis to see the distal end of the vagina visually.
  2. Place the tissue in 1x PBS to keep the tissue hydrated (Figure 5J).

Figure 5
Figure 5: Bladder, rectum, and cervix/vagina dissections. (A) Holding the bladder at an angle. (B) Cutting off the bladder. (C) Cutting the tendon connecting the cervix and rectum. (D) The tendon at 1.0x magnification. (E) Cutting the rectum. (F) Holding onto the cervix with forceps. (G) Cutting at the distal end of the vagina. (H) The bladder in a 35 mm Petri dish. (I) The rectum in a 35 mm Petri dish. (J) The cervix-vagina tissue complex in a 35 mm Petri dish. Please click here to view a larger version of this figure.

6. Sample preparation for tissue characterization

  1. Mechanical and visual analyses of the USL
    1. Place the USL with the pelvic attachment over a T-shaped wall within a custom staining well to ensure full immersion in the staining solution (CAD drawings of the well can be found in Supplementary Coding File 1 and Supplementary Coding File 2).
      NOTE: Use suture and forceps to help with the placement.
    2. Dilute a commercially available dye that stains free amine groups (5 µL, see Table of Materials) in 2.5 mL of 1x PBS, add the solution to the custom staining well, and stain the tissue for 2 h on a rocker at 4 °C.
      NOTE: Vortex the solution before adding it to the staining well.
    3. During the last 15 min of staining, add 2.5 µL of a commercially available dead cell nuclei stain (see Table of Materials) to the solution.
      NOTE: Vortex the solution prior to adding to the staining well.
  2. Raman analysis of the USLs
    1. Pin the USL in a straight line on a polydimethylsiloxane (PDMS) block that is contained in a custom well.
      NOTE: The PDMS is used as a soft substrate to enable the pinning of the sample in the desired configuration. Blocks of varying dimensions can be made by mixing the two components following the manufacturer's instructions (see Table of Materials), casting in a Petri dish, and, after polymerization, cutting the PDMS into the required geometry with a scalpel blade.
    2. Pin with insect pins at the suture loop and at the pelvic muscle. Hydrate the tissue with 1x PBS.
  3. Remaining tissues
    1. Snap-freeze the remaining tissues with liquid nitrogen or in an appropriate embedding compound, depending on the desired analyses.
    2. Save the tissues at −80 °C until subsequent analyses (e.g., immunohistochemical or biochemical assays).

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Representative Results

Each step of the dissection of a wild-type mouse is detailed in the associated video and figures related to the protocol. For this study, 6 week old female C57BL/6J mice were used (Supplementary Table 1). Three sample groups with USLs treated with different enzymes were analyzed: control (no treatment), collagenase-treated, and chondroitinase-treated groups. The smooth muscle, nerves, and lymphatics in the USL are surrounded by an ECM rich in fibrillar collagens and glycosaminoglycans (GAGs)5, which are thought to provide mechanical integrity to the tissue. The enzyme treatments were chosen to disrupt these ECM components and demonstrate the feasibility of resolving the biochemical and mechanical differences using Raman spectroscopy and mechanical (tensile) testing.

For digestion, all the USLs were placed in a thermomixer for 3 h in a solution of 1 mL at 37 °C and 300 rpm. The control USLs were placed in a solution of 1x PBS, the collagenase-treated USLs were placed in a solution of 1.0 U/mL collagenase type I, and the chondroitinase-treated USLs were placed in a solution of 2.0 U/mL chondroitinase ABC (see Table of Materials).

The USLs for mechanical testing were prepared based on step 6.1. The samples were placed in a custom loading chamber (a CAD drawing of the chamber can be found in Supplementary Coding File 3) for the mechanical testing protocol12. The pelvis was fixed in a stationary configuration, and the cervical end of the USL was attached to a buoy actuator, which was based on the design described in Jimenez et al.12 (Figure 6B; Supplementary Coding File 4), under an upright confocal microscope and attached to a 100 mN load cell (Figure 6A). The buoy system reduces the noise caused by the arm of the actuator when measuring force12. A pre-load of approximately 500 µN was used to set the reference configuration of the USL, and each USL was imaged while in this position. Then, a global displacement of 750 µm was applied and held for 5 min before imaging again. The USL was then brought back to the reference configuration, and this process was repeated for a 1,000 µm global displacement. This protocol was followed for all three sample groups (n = 1/group) to obtain the stress relaxation curves (Figure 7), and the peak and equilibrium stresses withstood by each USL were determined (Table 1). The stress was computed by dividing the measured force by the average cross-sectional area (Supplementary Table 2 and Supplementary Table 3).

Figure 6
Figure 6: Mechanical testing setup. (A) Loading chamber under the confocal microscope and connected to the actuator. (B) USL in the loading chamber; the pelvis remains stationary, while the cervical end is tied to an interconnected buoy system. Please click here to view a larger version of this figure.

Figure 7
Figure 7: Macroscopic axial stress-relaxation plot of murine USLs subjected to different enzyme treatments. The enzyme treatment appeared to reduce the average axial peak and relaxed stresses that the USL experiences at prescribed global displacements. (*) indicates relaxed stress values, which were measured 200 s after peak stress. Please click here to view a larger version of this figure.

Sample Global displacement (μm) Peak stress (kPa) Relaxed equilibrium stress (kPa)
Control 750 95 67
1000 135 97
Collagenase- treated 750 60 27
1000 94 60
Chondroitinase-treated 750 17 10
1000 25 16

Table 1: The effect of enzyme treatment on stress measurements of the USLs.

From the images taken (Figure 8), the strain fields were estimated using a combination of manual and automatic ("imregdemons" function in MATLAB) texture tracking of the ligament between the reference (unloaded) and deformed states13. Manual tracking was done by selecting the same cell nuclei in the reference and deformed images. Green-Lagrange strain fields were computed from the estimated displacement fields (Figure 9). The average axial strains (E11) were calculated for the intermediate portion of each specimen (Table 2).

Figure 8
Figure 8: Images of deformed and reference states of a control (untreated) specimen after global displacements. (A) A 750 µm global displacement. (B) A 1,000 µm global displacement. Green represents the reference state, and purple is the deformed state (scale bar = 500 µm). Please click here to view a larger version of this figure.

Figure 9
Figure 9: Strain fields estimation. The strain fields estimated for the samples in each treatment group demonstrate that the axial (E11), transverse (E22), and shear (E12) strains were spatially inhomogeneous, and the axial strain increased with increasing applied displacement (δ). Please click here to view a larger version of this figure.

Global Displacement (µm) Control Collagenase-treated  Chondroitinase-treated 
750 9.57% 8.01% 6.67%
1000 11.20% 15.75% 10.29%

Table 2: Average axial strains. The average axial strains for each mechanically tested specimen indicate that the global axial strain in the enzyme-treated samples was similar to or larger than in the control sample, suggesting that the reduction in macroscopic stress was due to enzyme treatment rather than smaller strains.

Confocal Raman spectroscopy (785 nm laser, 1.06 µm spot size) was conducted on the USLs prepared as described in steps 6.2 and following O'Brien et al.14 to semi-quantitatively evaluate the tissue biochemistry. Cosmic rays were identified and removed, the linear baseline was subtracted, and the intensity was normalized for each spectrum. The same treatment groups were compared (n = 3/group). Representative spectra for the intermediate section of each USL (Figure 10), as well as cervical and sacral ends (Supplementary Figure 1 and Supplementary Figure 2), are shown.

Figure 10
Figure 10: Influence of enzyme treatment on USL composition. Raman spectroscopy of the USL intermediate section suggests that enzyme treatments altered the biochemical composition of the murine USL. Please click here to view a larger version of this figure.

The peaks were correlated with different biological components based on the in vivo Raman spectroscopy of the human cervix performed by O'Brien et al.15. Our data were normalized using the peak of phosphatidylethanolamine (1,769 nm), a phospholipid found in cell membranes, which should remain unchanged after enzymatic treatments. Comparing across representative spectra revealed that the water content, collagens, hyaluronic acid, proteoglycans, and GAGs were decreased in both the collagenase-treated and chondroitinase-treated USLs (Figure 10, Supplementary Figure 1, and Supplementary Figure 2).

Supplementary Figure 1: Influence of enzyme treatment on USL composition. Raman spectroscopy of the USL cervical section suggests that the enzyme treatments altered the biochemical composition of the murine USL. Please click here to download this File.

Supplementary Figure 2: Influence of enzyme treatment on USL composition. Raman spectroscopy of the USL sacral section suggests that the enzyme treatments altered the biochemical composition of the murine USL. Please click here to download this File.

Supplementary Table 1: The weight, age, and USL analysis of the mice used in the study. Please click here to download this File.

Supplementary Table 2: Influence of enzyme treatment on USL force measurements. Please click here to download this File.

Supplementary Table 3: Cross-sectional area calculations of the mechanically tested USLs. Please click here to download this File.

Supplementary Coding File 1: CAD file for staining the well. Please click here to download this File.

Supplementary Coding File 2: CAD file for staining the well lid. Please click here to download this File.

Supplementary Coding File 3: CAD file of the loading chamber. Please click here to download this File.

Supplementary Coding File 4: CAD file of the buoy system. Please click here to download this File.

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The effect of structural damage on female reproductive tissues is understudied, and an easily accessible animal model for POP research is needed. The mouse is a cost-effective model that can mimic human reproductive studies16. Due to the growing interest in the study of the female reproductive system, there is a need for methods that aid the study of these tissues. To address this need, in this work, a method is established to dissect and prepare murine pelvic floor tissues for structural and functional analyses.

For the success of the dissection, adequate time and care are needed. Murine reproductive tissues are small and fragile. Attention to detail is required, especially while clearing the fat surrounding the USL and pelvic organs, to avoid damaging the USL or other tissues when the fat is removed. It is important to use the dissecting microscope to aid in identifying differences between the USL and fat, which the untrained eye can easily confuse. Removing the ureters can be helpful to avoid confusion with the USL, as the insertion points to the bladder and cervix are close to each other. When tying the suture around the USL, care must be taken to avoid puncturing any tissue by using forceps to gently lift the USLs to make space for the needle. To extract the USLs, the identification of the right tissue structure and key anatomical landmarks can be difficult, but with this method and practice, the dissection outcomes will be repeatable.

The proposed method outlines a detailed procedure for dissecting pelvic floor tissues from a single specimen. One limitation is that it is not possible to keep all the tissues completely intact since a piece of the cervix remains attached to the dissected USL to act as an anchor during the mechanical testing and help identify the orientation. Another limitation is that the mouse anatomy and human anatomy have differences in tissue shape, size, and orientation9. POP has been investigated using several animal models, such as rodents17,18, rabbits19,20 sheep20,21, swine5,22,23,24, and non-human primates25,26, as well as in human cadavers24,26,27, and some of these models have focused on the role of the USL. While each of these models is beneficial to the study of POP, an additional benefit of the mouse is the ease of genetic manipulation for creating novel disease models, which is not possible in larger animals27.

In order to take advantage of the mouse as a model, this method is developed to aid in the successful dissection of the USLs and the preparation of the tissues for various analyses, including mechanical testing, Raman microscopy, immunohistochemistry, and biochemical assays. With this method, it is possible to extract the USL with its anatomical connection to identify its orientation, to tether the USL, and to mechanically test the USL in conditions similar to those found in vivo.

The mechanical integrity of the USL is derived from the ECM components, including collagens, proteoglycans, and GAGs. Type I collagen is a major tensile load-bearing protein, and damage to this ECM component is thought to contribute to USL failure and POP. In this procedure, two different enzyme treatments were compared to test how varying the ECM composition affects the mechanical response and composition of the USL. The mechanical analysis suggested that collagen and GAGs contributed to the axial stiffness of the USL, lowering the average axial peak and equilibrium stresses (Figure 7). The enzyme-treated USLs experienced a similar or greater average axial strain (Table 2). Raman spectroscopy demonstrated that the collagenase-treated USL had decreased collagen content, as well as decreased water and hyaluronic acid, while the chondroitinase-treated USL had less hyaluronic acid, as well as lower water and collagen content. For these results, the Raman spectroscopy validated that the components were removed due to digestion. This method is anticipated to make pelvic floor-based studies more accessible and increase the number of investigators focusing on this understudied area.

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The authors have nothing to disclose.


This work was supported by the CU Boulder Summer Underground Research Opportunities Program (UROP) grant (C.B.), the NSF Graduate Research Fellowship (L.S.), the Schmidt Science Fellowship (C.L.), the University of Colorado Research & Innovation Seed Grant Program (2020 award to V.F., S.C., and K.C.), and the Anschutz Boulder Nexus Seed Grant at the University of Colorado (to V.F. and K.C.). Special acknowledgment goes to Dr. Tyler Tuttle for help with the loading chamber design as well as the Calve lab members for helpful discussions.


Name Company Catalog Number Comments
11 Blade Fisher 3120030 Removable blade
1x PBS Fisher BP399-1 Diluted from 10x concentration
Chondroitinase ABC Sigma C3667-10UN Enzyme 
Collagenase Type I Worthington Biochemical LS004194 Enzyme 
Confocal Microscope Leica STELLARIS 5 Upright configuration
Dissection Microscope Leica S9E With camera
Dumont #5 Forceps Fisher NC9626652 Thin tip
Female C57BL/6J mice Jackson Laboratory strain #: 000664
FemtoTools Micromanipulator FemtoTools FT-RS1002 100 mN load cell
FST Curved Forceps Fisher NC9639443 Curved tip
FST Sharp 9 mm Scissors  Fisher NC9639443 Dissection scissors
Ghost Dye 780  Tonbo 13-0865-T500 Free amine stain
Kimwipes Fisher 06-666 Box of 50 wipes
OCT Tissue Tek 4583 Used for tissue preservation
PDMS Thermo Fisher 044764.AK Follow manufacturer's instructions
Petri Dishes 35 mm Fisher FB0875711A Used for dissected tissue
Polyglactin 5-0 Suture Veter.Sut VS385VL With needle
Renishaw InVia Raman Microscope Renishaw PN192(EN)-02-A With confocal objectives
Rocking Platform VWR 10127-876 2 tier platform
Surgical Gloves Fisher 52818 For dissection 
Sytox Thermo Fisher S11381 Nuclear stain 
T-pins Fisher S99385 For dissection 
Transfer Pipets Fisher 13-711-7M For dissection 
Underpads Fisher 22037950 To cover dissection pad



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Bastías, C. S., Savard, L. M., Eckstein, K. N., Connell, K., Luetkemeyer, C. M., Ferguson, V. L., Calve, S. Isolation and Characterization of the Murine Uterosacral Ligaments and Pelvic Floor Organs. J. Vis. Exp. (193), e65074, doi:10.3791/65074 (2023).More

Bastías, C. S., Savard, L. M., Eckstein, K. N., Connell, K., Luetkemeyer, C. M., Ferguson, V. L., Calve, S. Isolation and Characterization of the Murine Uterosacral Ligaments and Pelvic Floor Organs. J. Vis. Exp. (193), e65074, doi:10.3791/65074 (2023).

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