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 &#38; 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.

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
<ol>
	<li>Euthanize the animal following the institutionally approved method. 
	NOTE: The present study used CO2 inhalation in alignment with the American Veterinary Medical Association&#39;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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>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).</li>
	<li>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).</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>Clear all of the fat tissue from the pelvic floor (Figure 2L-N). 
	&#8203;NOTE: Use forceps to gently pull and clear fat off the organs and tissues of interest.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65074/65074fig01.jpg" /> 
Figure 1: A clean workspace with all the tools needed to perform the dissections. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65074/65074fig02.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. USL harvesting
<ol>
	<li>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.</li>
	<li>Cut the ureters away from the bladder connection (Figure 3D). 
	NOTE: This is to avoid confusion with the USL.</li>
	<li>Cut the colon as close to the cervix as possible (Figure 3E,F). 
	NOTE: Irrigate the tissues with 1x PBS to maintain hydration.</li>
	<li>Place the mouse along with the dissection pad under the dissecting scope to visualize the USLs (Figure 3G).</li>
	<li>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.</li>
	<li>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)</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65074/65074fig03.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65074/65074fig04.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Bladder harvesting
<ol>
	<li>After the fat is cleared, hold the bladder with the forceps, and gently lift it at an angle of approximately 40&#176; (Figure 5A).</li>
	<li>With the scissors, slice off the bladder from the distal side, right above the cervix (Figure 5B).</li>
	<li>Place the tissue in a bath with 1x PBS to keep the tissue hydrated (Figure 5H).</li>
</ol>
4. Rectum harvesting
<ol>
	<li>Once the USLs are disconnected from the cervix and the bladder is dissected, lift the cervix at an angle of approximately 40&#176; 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).</li>
	<li>Cut the pubic bone at the pubic symphysis using scissors. Gently widen the workspace to increase visual access to the tissue insertions.</li>
	<li>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).</li>
	<li>Place the tissue in 1x PBS to keep the tissue hydrated (Figure 5I).</li>
</ol>
5. Cervix-vagina complex harvesting
<ol>
	<li>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.</li>
	<li>Place the tissue in 1x PBS to keep the tissue hydrated (Figure 5J).</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65074/65074fig05.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Sample preparation for tissue characterization
<ol>
	<li>Mechanical and visual analyses of the USL
	<ol>
		<li>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.</li>
		<li>Dilute a commercially available dye that stains free amine groups (5 &#181;L, 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 &#176;C. 
		NOTE: Vortex the solution before adding it to the staining well.</li>
		<li>During the last 15 min of staining, add 2.5 &#181;L 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.</li>
	</ol>
	</li>
	<li>Raman analysis of the USLs
	<ol>
		<li>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&#39;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.</li>
		<li>Pin with insect pins at the suture loop and at the pelvic muscle. Hydrate the tissue with 1x PBS.</li>
	</ol>
	</li>
	<li>Remaining tissues
	<ol>
		<li>Snap-freeze the remaining tissues with liquid nitrogen or in an appropriate embedding compound, depending on the desired analyses.</li>
		<li>Save the tissues at &#8722;80 &#176;C until subsequent analyses (e.g., immunohistochemical or biochemical assays).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

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.

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.

<table><tbody><tr><td>11 Blade</td><td>Fisher</td><td>3120030</td><td>Removable blade</td></tr><tr><td>1x PBS</td><td>Fisher</td><td>BP399-1</td><td>Diluted from 10x concentration</td></tr><tr><td>Chondroitinase ABC</td><td>Sigma</td><td>C3667-10UN</td><td>Enzyme&amp;nbsp;</td></tr><tr><td>Collagenase Type I</td><td>Worthington Biochemical</td><td>LS004194</td><td>Enzyme&amp;nbsp;</td></tr><tr><td>Confocal Microscope</td><td>Leica</td><td>STELLARIS 5</td><td>Upright configuration</td></tr><tr><td>Dissection Microscope</td><td>Leica</td><td>S9E</td><td>With camera</td></tr><tr><td>Dumont #5 Forceps</td><td>Fisher</td><td>NC9626652</td><td>Thin tip</td></tr><tr><td>Female C57BL/6J mice</td><td>Jackson Laboratory</td><td>strain #: 000664</td><td></td></tr><tr><td>FemtoTools Micromanipulator</td><td>FemtoTools</td><td>FT-RS1002</td><td>100 mN load cell</td></tr><tr><td>FST Curved Forceps</td><td>Fisher</td><td>NC9639443</td><td>Curved tip</td></tr><tr><td>FST Sharp 9 mm Scissors&amp;nbsp;</td><td>Fisher</td><td>NC9639443</td><td>Dissection scissors</td></tr><tr><td>Ghost Dye 780&amp;nbsp;</td><td>Tonbo</td><td>13-0865-T500</td><td>Free amine stain</td></tr><tr><td>Kimwipes</td><td>Fisher</td><td>06-666</td><td>Box of 50 wipes</td></tr><tr><td>OCT</td><td>Tissue Tek</td><td>4583</td><td>Used for tissue preservation</td></tr><tr><td>PDMS</td><td>Thermo Fisher</td><td>044764.AK</td><td>Follow manufacturer&#039;s instructions</td></tr><tr><td>Petri Dishes 35 mm</td><td>Fisher</td><td>FB0875711A</td><td>Used for dissected tissue</td></tr><tr><td>Polyglactin 5-0 Suture</td><td>Veter.Sut</td><td>VS385VL</td><td>With needle</td></tr><tr><td>Renishaw InVia Raman Microscope</td><td>Renishaw</td><td>PN192(EN)-02-A</td><td>With confocal objectives</td></tr><tr><td>Rocking Platform</td><td>VWR</td><td>10127-876</td><td>2 tier platform</td></tr><tr><td>Surgical Gloves</td><td>Fisher</td><td>52818</td><td>For dissection&amp;nbsp;</td></tr><tr><td>Sytox</td><td>Thermo Fisher</td><td>S11381</td><td>Nuclear stain&amp;nbsp;</td></tr><tr><td>T-pins</td><td>Fisher</td><td>S99385</td><td>For dissection&amp;nbsp;</td></tr><tr><td>Transfer Pipets</td><td>Fisher</td><td>13-711-7M</td><td>For dissection&amp;nbsp;</td></tr><tr><td>Underpads</td><td>Fisher</td><td>22037950</td><td>To cover dissection pad</td></tr></tbody></table>

isolation and characterization of the murine uterosacral ligaments and pelvic floor organs

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.

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 &#176;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 &#181;N 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 &#181;m 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 &#181;m 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).
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65074/65074fig06.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/65074/65074fig07.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample</td>
			<td>Global displacement (&#956;m)</td>
			<td>Peak stress (kPa)</td>
			<td>Relaxed equilibrium stress (kPa)</td>
		</tr>
		<tr>
			<td rowspan="2">Control</td>
			<td>750</td>
			<td>95</td>
			<td>67</td>
		</tr>
		<tr>
			<td>1000</td>
			<td>135</td>
			<td>97</td>
		</tr>
		<tr>
			<td rowspan="2">Collagenase- treated</td>
			<td>750</td>
			<td>60</td>
			<td>27</td>
		</tr>
		<tr>
			<td>1000</td>
			<td>94</td>
			<td>60</td>
		</tr>
		<tr>
			<td rowspan="2">Chondroitinase-treated</td>
			<td>750</td>
			<td>17</td>
			<td>10</td>
		</tr>
		<tr>
			<td>1000</td>
			<td>25</td>
			<td>16</td>
		</tr>
	</tbody>
</table>
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 (&#34;imregdemons&#34; 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).
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/65074/65074fig08.jpg" /> 
Figure 8: Images of deformed and reference states of a control (untreated) specimen after global displacements. (A) A 750 &#181;m global displacement. (B) A 1,000 &#181;m global displacement. Green represents the reference state, and purple is the deformed state (scale bar = 500 &#181;m). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/65074/65074fig09.jpg" /> 
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 (&#948;). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Global Displacement (&#181;m)</td>
			<td>Control</td>
			<td>Collagenase-treated&#160;</td>
			<td>Chondroitinase-treated&#160;</td>
		</tr>
		<tr>
			<td>750</td>
			<td>9.57%</td>
			<td>8.01%</td>
			<td>6.67%</td>
		</tr>
		<tr>
			<td>1000</td>
			<td>11.20%</td>
			<td>15.75%</td>
			<td>10.29%</td>
		</tr>
	</tbody>
</table>
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 &#181;m spot size) was conducted on the USLs prepared as described in steps 6.2 and following O&#39;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.
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/65074/65074fig10v3.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074fig10v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The peaks were correlated with different biological components based on the in vivo Raman spectroscopy of the human cervix performed by O&#39;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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/2023-01-18 supplemental figure 1.pdf" target="_blank">Please click here to download this File.</a>
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/2023-01-18 supplemental figure 2.pdf" target="_blank">Please click here to download this File.</a>
Supplementary Table 1: The weight, age, and USL analysis of the mice used in the study. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074_Supplementary Table 1_rev.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 2: Influence of enzyme treatment on USL force measurements. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074_Supplementary Table 2_rev.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 3: Cross-sectional area calculations of the mechanically tested USLs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/65074_Supplementary Table 3_rev.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Coding File 1: CAD file for staining the well. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/S1 staining well lid.zip" target="_blank">Please click here to download this File.</a>
Supplementary Coding File 2: CAD file for staining the well lid. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/S2 staining well_groove.zip" target="_blank">Please click here to download this File.</a>
Supplementary Coding File 3: CAD file of the loading chamber. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/S3 loading chamber.zip" target="_blank">Please click here to download this File.</a>
Supplementary Coding File 4: CAD file of the buoy system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65074/S4 buoy_3.2.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Isolation and Characterization of the Murine Uterosacral Ligaments and Pelvic Floor Organs at JoVE.com

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

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 ...

isolation-characterization-murine-uterosacral-ligaments-pelvic-floor

Nanyang Technological University

Research

JoVE Journal

Biology

1.2K Views.  University of Colorado Boulder. 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.

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

Mouse Model of Surgically-induced Endometriosis by Auto-transplantation of Uterine Tissue

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of lesions, and/or chronic pharmaceutical management of pain and infertility symptoms. The cost associated with endometriosis has been estimated at 22 billion dollars per year in the United States1. To further our understanding of mechanisms underlying this enigmatic disease, animal models have been employed. Primates spontaneously develop endometriosis and therefore primate models most closely resemble the disease in women. Rodent models, however, are more cost effective and readily available2. The model that we describe here involves an autologous transfer of uterine tissue to the intestinal mesentery (Figure 1) and was first developed in the rat3 and later transferred to the mouse4. The goal of the autologous rodent model of surgically-induced endometriosis is to mimic the disease in women. We and others have previously shown that the altered gene expression pattern observed in endometriotic lesions from mice or rats mirrors that observed in women with the disease5,6. One advantage of performing the surgery in the mouse is that the abundance of transgenic mouse strains available can aid researchers in determining the role of specific components important in the establishment and growth of endometriosis. An alternative model in which excised human endometrial fragments are introduced to the peritoneum of immunocompromised mice is also widely used but is limited by the lack of a normal immune system which is thought to be important in endometriosis2,7. Importantly, the mouse model of surgically induced endometriosis is a versatile model that has been used to study how the immune system8, hormones9,10 and environmental factors11,12 affect endometriosis as well as the effects of endometriosis on fertility13 and pain14.

A description of the surgical induction of endometriosis in mice and rats by auto-transplantation of uterine tissue to the arterial cascade of the intestinal mesentery.

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of ...

Medicine

Formation of Human Prostate Epithelium Using Tissue Recombination of Rodent Urogenital Sinus Mesenchyme and Human Stem Cells

Progress in prostate cancer research is severely limited by the availability of human-derived and hormone-na&iuml;ve model systems, which limit our ability to understand genetic and molecular events underlying prostate disease initiation. Toward developing better model systems for studying human prostate carcinogenesis, we and others have taken advantage of the unique pro-prostatic inductive potential of embryonic rodent prostate stroma, termed urogenital sinus mesenchyme (UGSM). When recombined with certain pluripotent cell populations such as embryonic stem cells, UGSM induces the formation of normal human prostate epithelia in a testosterone-dependent manner. Such a human model system can be used to investigate and experimentally test the ability of candidate prostate cancer susceptibility genes at an accelerated pace compared to typical rodent transgenic studies. Since Human embryonic stem cells (hESCs) can be genetically modified in culture using inducible gene expression or siRNA knock-down vectors prior to tissue recombination, such a model facilitates testing the functional consequences of genes, or combinations of genes, which are thought to promote or prevent carcinogenesis.
The technique of isolating pure populations of UGSM cells, however, is challenging and learning often requires someone with previous expertise to personally teach. Moreover, inoculation of cell mixtures under the renal capsule of an immunocompromised host can be technically challenging. Here we outline and illustrate proper isolation of UGSM from rodent embryos and renal capsule implantation of tissue mixtures to form human prostate epithelium. Such an approach, at its current stage, requires in vivo xenografting of embryonic stem cells; future applications could potentially include in vitro gland formation or the use of induced pluripotent stem cell populations (iPSCs).

To unravel the earliest molecular mechanisms underlying prostate cancer initiation, novel and innovative human model systems and approaches are desperately needed. The potential of pre-prostatic urogenital sinus mesenchyme (UGSM) to induce pluripotent stem cell populations to form human prostate epithelium is a powerful experimental tool in prostate research.

Progress in prostate cancer research is severely  limited by the availability of human-derived and hormone-na&iuml;ve model systems,  which limit our ...

Assessment of Perigenital Sensitivity and Prostatic Mast Cell Activation in a Mouse Model of Neonatal Maternal Separation

Chronic prostatitis/chronic pelvic pain syndrome (CP/CPPS) has a lifetime prevalence of 14% and is the most common urological diagnosis for men under the age of 50, yet it is the least understood and studied chronic pelvic pain disorder. A significant subset of patients with chronic pelvic pain report having experienced early life stress or abuse, which can markedly affect the functioning and regulation of the hypothalamic-pituitary-adrenal (HPA) axis. Mast cell activation, which has been shown to be increased in both urine and expressed prostatic secretions of CP/CPPS patients, is partially regulated by downstream activation of the HPA axis. Neonatal maternal separation (NMS) has been used for over two decades to study the outcomes of early life stress in rodent models, including changes in the HPA axis and visceral sensitivity. Here we provide a detailed protocol for using NMS as a preclinical model of CP/CPPS in male C57BL/6 mice. We describe the methodology for performing NMS, assessing perigenital mechanical allodynia, and histological evidence of mast cell activation. We also provide evidence that early psychological stress can have long-lasting effects on the male urogenital system in mice.

We are measuring perigenital mechanical sensitivity and mast cell activation in the prostate of male C57BL/6 mice that underwent an early life stress paradigm &#8211; neonatal maternal separation, in order to induce a preclinical model of chronic prostatitis/chronic pelvic pain syndrome.

Chronic prostatitis/chronic pelvic pain syndrome (CP/CPPS) has a lifetime prevalence of 14% and is the most common urological diagnosis for men under the ...

Transvaginal Mesh Insertion in the Ovine Model

This protocol describes mesh insertion into the rectovaginal septum in sheep using a single vaginal incision technique, with and without the trocar-guided insertion of anchoring arms. Parous sheep underwent the dissection of the rectovaginal septum, followed by the insertion of an implant with or without four anchoring arms, both designed to fit the ovine anatomy. The anchoring arms were put in place using a trocar and an "outside-in" technique. The cranial arms were passed through the obturator, gracilis, and adductor magnus muscles. The caudal arms were fixed near the sacrotuberous ligament, through the coccygeus muscles. This technique allows for the mimicking of surgical procedures performed in women suffering from pelvic organ prolapse. The anatomical spaces and elements are easily identified. The most critical part of the procedure is the insertion of the cranial trocar, which can easily penetrate the peritoneal cavity or the surrounding pelvic organs. This can be avoided by a more extensive retroperitoneal dissection and by guiding the trocar more laterally. This approach is designed only for experimental testing of novel implants in large animal models, as trocar-guided insertion is currently not used clinically.

This protocol describes mesh implantation in the ovine rectovaginal septum using a single vaginal incision technique, with and without the trocar-guided insertion of anchoring arms.

This protocol describes mesh insertion into the rectovaginal septum in sheep using a single vaginal incision technique, with and without the trocar-guided ...

Dissection of Pelvic Autonomic Ganglia and Associated Nerves in Male and Female Rats

The bilateral major pelvic ganglia (MPG; synonym, pelvic ganglia) are the primary source of postganglionic sympathetic and parasympathetic neurons innervating pelvic organs of rodents; the functionally equivalent structure in humans is the inferior hypogastric plexus. The major pelvic ganglia also provide the route by which lumbar and sacral sensory axons reach the pelvic organs. These complex, mixed ganglia can prove challenging to identify and dissect for further experimental study of normal autonomic mechanisms or to establish preclinical models of disease, injury or visceral pain. Here we describe a protocol to access and visualize these ganglia and their associated nerve tracts. We provide this protocol with schematics for both male and female rats, as the ganglion size and landmarks for identification differ between sexes. The protocol describes removal of the ganglion for in vitro studies, but this method can be integrated into a surgical recovery protocol for experimental interventions (e.g., nerve crush, nerve resection) or for mapping neuronal circuits (e.g., by microinjection of neural tracers). We also demonstrate the primary structures of the ganglion and its associated nerves immediately following dissection and following immunohistochemical staining.

The major pelvic ganglia contain parasympathetic and sympathetic neurons that innervate pelvic organs. Here we describe a dissection method and provide schematics for identification of these ganglia and their associated nerves. These methods can be applied to experimental manipulation of these ganglia in vivo or removal post-mortem for further study.

The bilateral major pelvic ganglia (MPG; synonym, pelvic ganglia) are the primary source of postganglionic sympathetic and parasympathetic neurons ...

Neuroscience

Robot-assisted Total Mesorectal Excision and Lateral Pelvic Lymph Node Dissection for Locally Advanced Middle-low Rectal Cancer

Since their approval for clinical use, da Vinci surgical robots have shown great advantages in gastrointestinal surgical operations, especially in complex procedures. The high-quality 3-D visual, multijoint arm and natural tremor filtration allow the surgeon to expose and dissect more accurately with minimal invasion. Total mesorectal excision is the standard surgical technique for the treatment of resectable rectal cancer. To reduce the lateral recurrence rate, lateral pelvic lymph node dissection can be performed, as it is a safe and feasible procedure for locally advanced middle-low rectal cancer with a high possibility of metastasis to the lateral lymph nodes. However, the complexity of the anatomic structures and the high postoperative complication rate limit its application. Recently, several surgeons have increasingly used robotic techniques for total mesorectal excision and lateral pelvic lymph node dissection. Compared with open and laparoscopic surgery, the robotic technique has several advantages, such as less blood loss, fewer blood transfusions, minimal trauma, shorter postoperative hospitalization, and quicker recovery. A robotic approach is generally regarded as a reasonable alternative for complicated procedures such as lateral pelvic lymph node dissection, although there are a limited number of high-quality prospective randomized controlled studies reporting direct evidence. Here, we provide the detailed steps of robot-assisted total mesorectal excision and lateral pelvic lymph node dissection performed at the First Affiliated Hospital of Xi&#39;an Jiaotong University.

The robotic technique described herein aims to detail a stepwise approach to robot-assisted total mesorectal excision and lateral pelvic lymph node dissection for locally advanced (T3/T4) rectal cancer located below the peritoneal reflection.

Since their approval for clinical use, da Vinci surgical robots have shown great advantages in gastrointestinal surgical operations, especially in complex ...

Development of a Uterosacral Ligament Suspension Rat Model

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.

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.

Pelvic organ prolapse (POP) is a common pelvic floor disorder (PFD) with the potential to significantly impact a woman's quality of life. Approximately ...

Laparoscopic Non-Mesh Cerclage Pectopexy for Pelvic Organ Prolapse

Pelvic organ prolapse (POP) is widespread among the female population and significantly impairs the patient's quality of life. It is important to restore apical support for treating POP. Sacrocolpopexy and pectopexy are indicated for apical prolapse. Using a synthetic mesh in these techniques increases success by enhancing apical support. However, the implantation of synthetic mesh is associated with mesh-related complications. In addition, the exorbitant cost of synthetic mesh and lack of universal access limit the popularity of these procedures. The current study develops a unique technique known as laparoscopic non-mesh cerclage pectopexy (LNMCP), in which permanent cervical cerclage sutures are embedded in the round ligament until the iliopectineal ligament. The iliopectineal ligament was sutured, resulting in a firm cervical suspension. The procedure was successfully performed in 16 cases in the hospital. The surgical duration was 67.8 min &plusmn; 15.5 min, and the blood loss was 73.1 mL &plusmn; 51.1 mL. No procedural complications were seen. LNMCP is associated with an objective success rate of 100% and a subjective success rate of 93.8%. LNMCP for patients with apical prolapse obviates the need for a mesh, thereby avoiding complications associated with mesh erosion and reducing medical costs. In addition, it is easy to perform even in resource-poor areas without access to synthetic mesh.

The present pilot study describes the development of laparoscopic non-mesh cerclage pectopexy to treat pelvic organ prolapse. The procedure can be used to prevent any complications associated with the use of mesh.

Pelvic organ prolapse (POP) is widespread among the female population and significantly impairs the patient's quality of life. It is important to restore ...

Targeting Mouse Sympathetic Neurons: A Lumbar Sympathectomy Approach

Surgical Lumbar Sympathectomy in Mice

Peripheral nerve injuries are common, and full functional recovery after injury is achieved in only 10% of patients. The sympathetic nervous system plays many critical roles in maintaining bodily homeostasis, but it has rarely been studied in the context of peripheral nerve injury. The extent of postganglionic sympathetic neuronal functions in distal targets in the periphery is currently unclear. To better explore the role of sympathetic innervation of peripheral targets, a surgical &#34;knock-out&#34; model provides an alternative approach. Although this can be achieved chemically, chemical destruction of postganglionic sympathetic neurons can be nonspecific and dose-dependent. The use of a surgical lumbar sympathectomy in mice, once thought to be &#34;virtually not practicable&#34; in small animals, allows for specific targeting of postganglionic sympathetic neurons that innervate the hind limbs. This manuscript describes how to surgically remove the L2-L5 lumbar sympathetic ganglia from a mouse as a survival surgery, which reliably decreases the hind paw sweat response and the number of sympathetic axons in the sciatic nerve.

Author Spotlight: Exploring Plasticity of Sympathetic Neurons

This manuscript presents a protocol for surgically removing the postganglionic lumbar sympathetic neurons from a mouse. This procedure will facilitate a multitude of studies aimed at investigating the role of sympathetic innervation in distal tissue targets.

Peripheral nerve injuries are common, and full functional recovery after injury is achieved in only 10% of patients. The sympathetic nervous system plays ...

Ex Vivo Method for Assessing the Mouse Reproductive Tract Spontaneous Motility and a MATLAB-based Uterus Motion Tracking Algorithm for Data Analysis

Dysmenorrhea, or painful cramping, is the most common symptom associated with menses in females and its severity can hinder women&#39;s everyday lives. Here, we present an easy and inexpensive method that would be instrumental for testing new drugs decreasing uterine contractility. This method utilizes the unique ability of the entire mouse reproductive tract to exhibit spontaneous motility when maintained ex vivo in a Petri dish containing oxygenated Krebs buffer. This spontaneous motility resembles the wave-like myometrial activity of the human uterus, referred to as endometrial waves. To demonstrate the effectiveness of the method, we employed a well-known uterine relaxant drug, epinephrine. We demonstrate that the spontaneous motility of the entire mouse reproductive tract can be quickly and reversibly inhibited by 1 &#181;M epinephrine in this Petri dish model. Documenting the changes of uterine motility can be easily done using an ordinary smart phone or a sophisticated digital camera. We developed a MATLAB-based algorithm allowing motion tracking to quantify spontaneous uterine motility changes by measuring the rate of uterine horn movements. A major advantage of this ex vivo approach is that the reproductive tract remains intact throughout the entire experiment, preserving all intrinsic intrauterine cellular interactions. The major limitation of this approach is that up to 10-20% of uteri may exhibit no spontaneous motility. Thus far, this is the first quantitative ex vivo method for assessing spontaneous uterine motility in a Petri dish model.

Uterine contractions are important for the well-being of females. However, pathologically increased contractility may result in dysmenorrhea, especially in younger females. Here, we describe a simple ex vivo preparation allowing quick assessment of the efficacy of smooth muscle relaxants that may be used for treating dysmenorrhea.

Dysmenorrhea, or painful cramping, is the most common symptom associated with menses in females and its severity can hinder women&#39;s everyday lives. ...