We thank Ivan Laermans, Rosita Kinart, Ann Lissens (Centre for Surgical Technologies, KU Leuven, Leuven, Belgium). Jo Verbinnen and Kristof Reyniers (Vesalius Institute of Anatomy, Faculty of Medicine, KU Leuven, Leuven, Belgium) provided technical support during the experiment. We thank Leen Mortier for the help with data and manuscript management. We thank FEG Textiltechniken for manufacturing prototype meshes, sterilizing them, and donating them unconditionally for research.

Ethical approval for this experiment was obtained from the Ethics Committee on Animal Experimentation of the KU Leuven (P065/2013). Animals were treated in accordance with current national guidelines on animal welfare.
1. Material and the Experimental Animal
<ol>
	<li>Surgery preparation
	<ol>
		<li>In the surgical theater cover a table with a sterile drape and prepare one sterile curved trocar (Figure 1, panel A), sterile surgical instruments, sutures, and sterile gauze. Perform the entire surgical procedure in sterile conditions if the experiment includes follow-up. Place all instruments on the table to be ready for use during surgery.</li>
		<li>Remove a sterile rectangular implant and/or an implant with anchoring arms from the sterile package and put them on the table covered with sterile drape (Figure 1B, C, and D).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/55706/55706fig1.jpg" /> 
Figure 1: Trocar and Implants. (A) Schematic drawing of the trocar. (B) H-shaped polyvinylidene fluoride (PVDF) implant, with a detail of the central part (panel C). Its shape was inspired by the four-arm meshes currently available for transvaginal prolapse repair. The rectangular body (30 x 40 mm2) is laterally extended by four outstretched arms (150 x 10 mm2). The dimensions of the arms are designed to be long enough to pierce the relevant suspension structures, based on earlier anatomical studies17. (D) The rectangular implant (30 x 40 mm2). Both implants were made of polyvinylidene fluoride; textile characteristics and properties are in Table 1.
<ol start="2">
	<li>Experimental animal (Ewe, 45-60 kg)
	<ol>
		<li>Administer premedication of 1 mL of 15 mg/mL atropine sulfate and 1 mL/50 kg of xylazine HCl intramuscularly (i.m.) 30 min before the surgical procedure.</li>
		<li>After 30 min, ensure that the premedication has made the sheep lethargic and sleepy.</li>
		<li>Insert an intravenous catheter into the jugular vein and administer 0.075 mL/kg of ketamine 100 mg/mL HCl. Confirm deep anesthesia by observing the lack of reaction to painful stimuli.</li>
		<li>Move the animal onto the surgical table and secure its airways by intubation. Maintain the anesthesia with 2.5% isoflurane in 5 L/min oxygen.</li>
		<li>Keep the intravenous line inserted in the jugular vein and supply 500 mL of saline solution at a flow rate of 150 mL/h.</li>
		<li>Administer prophylactic antibiotics i.m. (amoxicillin clavulanate, 7 mg/kg) and post-operative analgesics (buprenorfin and chlorocresol, 1 mL) or the equivalent according to local protocols.</li>
		<li>Place the animal in lithotomy position on the end of the surgical table and secure its limbs, with the hips in hyper-flexion, using ropes (Figure 2, panel A).</li>
		<li>Manually empty the bladder and rectum by pushing on them trans-vaginally.</li>
		<li>Shave the perineum, the medial part of the thigh, and the tail folds and disinfect with polyvidone iodium 7.5% (Figure 2, panel B and C).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/55706/55706fig2.jpg" /> 
Figure 2: Animal Surgery. (A) A sheep placed in the supine position, with the hips hyper-flexed by securing the lower limbs. (B) The external entrance points for trocar insertion are on the ventral side (empty arrow) and dorsally on the lateral tail folds (full arrow). (C) Position of the ventral insertion points; the dashed line in the middle represents the midsagittal plane of the animal. (D) Dissected rectovaginal septum. (E) Insertion of the ventral trocar through the muscles on the medial side of the thigh, the obturator foramen, and the paravaginal space. The trajectory of the piercing trocar is controlled with the finger. (F and G) Once the trocar is in place, the wire sling (open arrow) is advanced and loaded with the arm of the vaginal mesh. (H) Final position of the ventral (full arrows) and dorsal (empty arrow) arms. (I) The central part is placed tension-free between the vaginal wall and the rectal adventitia.
<ol start="10" style="margin-left: 40px;">
	<li>Prepare personnel for a surgery in sterile conditions. Put on a surgical cap and mouth mask, wash hands for surgery, and put on a surgical gown and sterile gloves.</li>
	<li>Cover the animal with a sterile drape and make an opening above the genital hiatus.</li>
</ol>
2. Surgical Procedure
<ol>
	<li>Preparation of the rectovaginal septum
	<ol>
		<li>Grasp the dorsal vaginal wall 3 cm cranial to the hymeneal ring using Allis forceps.</li>
		<li>Take a syringe loaded with 10 mL of saline and fitted with a 22 G needle. Insert it through the vaginal epithelium (approximately 3 - 4 mm deep) and into the midline of the rectovaginal septum, 1.5 cm cranial to the hymeneal ring.</li>
		<li>Perform &#34;aqua-dissection&#34; by injecting saline in the rectovaginal septum11.</li>
		<li>Make a 3 cm-long midline incision on the vaginal epithelium, starting caudal to the Allis forceps (step 2.1.1) and ending at the hymeneal ring using a scalpel. Enter the recto-vaginal space through this incision.</li>
		<li>Place the self-retaining retractor (see the Table of Materials) over the perineum and place four sharp stay hooks in the vaginal incision to keep it open.</li>
		<li>With your finger, bluntly dissect the recto-vaginal fascia from the vaginal wall laterally towards the pelvic side walls and cranially up to the caudal aspect of the cul-de-sac. Create suitable space for the 30 x 40 mm2 central part of the mesh (Figure 2, panel D).</li>
		<li>Perform haemostasis with haemostatic forceps or a crisscross haemostatic ligature whenever necessary. 
		NOTE: Small bleeders can be clamped with the hemostatic forceps. This crushes the vessel and initiates the natural coagulation cascade. For stronger bleeding, grasp the bleeding vessel with forceps and place a crisscross ligature, securing it with a square knot. At this point, one can either insert the rectangular implant (step 2.2) or continue with the dissection to insert the implant with anchoring arms (step 2.3).</li>
	</ol>
	</li>
	<li>Flat mesh insertion
	<ol>
		<li>Insert the vaginal retractor into the vaginal incision to allow a better view of the cranial part of the dissected area.</li>
		<li>Suture the left and right cranial corner of the implant with a simple interrupted 3/0 polypropylene suture on the left and right sides of the most cranial aspect of the dissected recto-vaginal space. Cut the residual suture material. Keep the suture away from the vaginal lumen (i.e., do not penetrate the vaginal wall). 
		NOTE: The implant is always sutured to the connective tissue comprising the recto-vaginal septum. The vaginal wall is not penetrated if the suture material cannot be seen in the vagina.</li>
		<li>Add one additional simple interrupted suture midway along the cranial aspect of the implant.</li>
		<li>Suture the lateral edges of the implant midway onto the surrounding connective tissue with a simple interrupted 3/0 polypropylene. Keep the implant as flat as possible and tension-free.</li>
		<li>Suture the left and right caudal corners with simple interrupted 3/0 polypropylene sutures on the left and right sides of the most caudal aspect of the rectovaginal space.</li>
		<li>Add one additional simple interrupted suture midway along the caudal aspect of the implant.</li>
		<li>Close the vaginal incisions with a running 3/0 polyglactin suture.</li>
	</ol>
	</li>
	<li>Insertion and anchoring of implant with arms (trocar-guided technique).
	<ol>
		<li>Continue the dissection of the recto-vaginal space created in step 2.1 cranio-ventrally to reach the medial aspect of the obturator foramen, which can easily be palpated.</li>
		<li>Dissect the space caudo-laterally to reach the caudal aspect of the sacrotuberous ligament and the caudally located coccygeus muscle.</li>
		<li>With a no. 24 blade, make four 1 cm-wide incisions on the vulvar side, cutting through the skin and superficial muscular fascia (Figure 2, panel B and C).</li>
		<li>Make two &#8220;ventral&#8221; incisions on the medial aspect of the thigh, proximately 4 cm cranial from the caudal border of the sciatic arch (i.e., the inferior border of the symphysis) and 3 cm lateral from the midline (Figure 2, panel C).</li>
		<li>Make two &#34;dorsal&#34; incisions at the level of the insertion of the tail folds, 2 cm medial to the tuber ischiadicum, which can be easily palpated (Figure 2, panel B).</li>
		<li>Place a curved trocar through one of the ventral incisions (Figure 2, panel E).</li>
		<li>Pass the trocar through the adductor magnus muscle, the external obturator, and the medial aspect of the obturator foramen.</li>
		<li>Control the progression of the trocar with a finger inserted through the vaginal incision. Guide its tip to the tendinous arc of the levator ani muscle (Figure 2, panel E).</li>
		<li>Expose the guiding wire in the vaginal wall incision and load it with the corresponding ipsilateral cranial mesh arm (Figure 2, panel F).</li>
		<li>Pull the trocar loaded with the mesh arm through the above structures. Keep the arm tension-free.</li>
		<li>Repeat the process with the second cranial arm through the ventral incision on the other side of the animal.</li>
		<li>Through one of dorsal incisions, pass the trocar through the coccygeus muscle, just distal to the sacrotuberous ligament (Figure 2, panel G).</li>
		<li>Expose the guide wire through the vaginal incision, grasp the dorsal arm of the mesh, and pull it out. Keep the arm tension-free and repeat on the other side.</li>
		<li>Adjust the position of the mesh by flattening it and applying tension to the arms, but keep the mesh tension-free (Figure 2, panel I; Figure 3).</li>
		<li>Fix the body of the mesh with a simple interrupted 3/0 polypropylene suture in the middle of its caudal border, securing it to the surrounding connective tissue.</li>
		<li>Cut the arms at the level of the skin and close all skin incisions with simple interrupted 3/0 polyglecaprone sutures (Figure 2, panel H).</li>
		<li>Close the vaginal incision with a running 3/0 polyglecaprone suture.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" src="/files/ftp_upload/55706/55706fig3.jpg" /> 
Figure 3: Schematic Illustration of the Ovine Pelvis, with the Cranial Arms Passing through the Obturator Foramen and the Caudal Arms Passing through the Tail Folds. The broad sacrotuberous ligament is in blue. The smaller panel illustrates the position of the arms on an animal in recumbent position, just before shortening the excessive amount of material. The main panel shows the same but with the skin and muscles removed.

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This research program on the ovine model was supported by an unconditional grant from Medri and Blasingame, Burch, Garrard and Ashley (Atlanta GA, USA). Agreements are handled via the Leuven Research and Development transfer office. Sponsors did not interfere with the planning, execution, or reporting of this experiment, nor do they own the results. NS and LH are recipients of a grant from the EC in the FP7-framework (Bip-Upy project; NMP3-LA-2012-310389). AF was supported by a grant from the EC in the industry-academic partnership program (251356).

Here, we describe an experimental procedure in sheep, aimed to mimic vaginal dissection and transvaginal mesh insertion of an implant with or without anchoring arms. The subsequent steps and instruments were inspired by surgical procedures done for POP and stress urinary incontinence15,16,19,20. After initial anatomical dissections, there were still some problems during experimental mesh insertion. In the first animal, a perforation in the peritoneal cavity at the level of de cul-de-sac was found. This has been previously described clinically in women21. In subsequent procedures, this was avoided by guiding the tip of the trocar more laterally (i.e., closer to the pelvic side wall). In later sheep, no other complications were observed.
The most feared anatomical structures were the obturator nerve and artery. In humans, the distance between the trocar/implant and the obturator canal is 1.9 - 3.0 cm22. The high variability of the trocar/implant position in women was previously explained by the exact positioning of the legs or the needle trajectory. Therefore, the hind limbs of the sheep were secured in hyper-flexion at the hips to allow greater access to the vagina. In this position the gracilis muscle is moved cranially. As a consequence, the trocar passed through the adductor magnus muscle, which is undivided but well-developed in sheep23. A more cranial passage of the trocar is possible yet may harm structures in the obturator canal.
Similarly to what is described clinically, the arms fixed the implant to given anatomical locations, corresponding to natural attachments of the rectovaginal septum in sheep23. The implant lay flat, apparently supporting the posterior vaginal wall without extending more laterally. Consequently, it did not have a tendency to fold. It may be possible to use larger implants. However, as demonstrated before in sheep, larger implants are associated with an up to 50% retraction and more local graft-related complications9.
Previous investigation of novel implants in various animal models included the sheep as a model for vaginal surgery. One aim of this work was to fix implants in a manner similar to that done clinically (i.e., by transfixing the arms of the mesh to anatomical structures in the pelvis). This procedure was not described earlier in a sheep model. This surgery follows a strategy similar to what was used to introduce new needle- and trocar-assisted surgeries into clinical practice15. However, these findings seem to be less relevant than a few years ago, as the use of vaginal implants and thus, trocar-guided procedures are quickly dropping due to the consecutive health warnings by the FDA and SCENIHR24,25.
Though we describe herein the technique, the number of animals in this experiment was very limited, so the full level of anatomical variability may not be represented here. Another limitation is that this technique describes a posterior compartment procedure, which may be less practiced. There are a few reports on surgery in the anterior compartment in sheep13, but smaller implants were usually used and complications were more frequent. Though these findings may be sufficient to plan further experiments, the feasibility of anterior vaginal mesh placement may also have been informative.
In conclusion, this is a description of a safe and feasible surgical technique in the ovine animal model for vaginal surgery that permits trans-vaginal and trocar-guided tension-free vaginal implant insertion. Relevant comparable structures could be blindly pierced without obvious risks for vessel, nerve, or organ injury. This model can of course also be used for simulated vaginal surgery using native tissue.

Pelvic organ prolapse is clinically diagnosed in half of women who had at least one vaginal delivery, but subjectively, it bothers half of women overall1. The mainstay of therapy is surgical reconstruction using either native tissue or implant materials, but each of these methods has its limitations, including recurrence or local complications2,3,4. The ideal implant has not yet been identified; hence, there is an ongoing demand for product innovation and for the development of a proper pipeline for preclinical experimentation prior to the introduction of new products and techniques to the market. One of the steps in this track is experimental evaluation on suitable animal models5,6. Ideally, they should mimic the anatomical, biomechanical, and biological environments. When it comes to the experimental evaluation of novel implants, they are typically tested first in smaller models, either for biocompatibility or for the reconstruction of abdominal wall defects. That type of experiments has been criticized, because the implants are not inserted into the area of interest (i.e., the vagina)7. Vaginal surgery models are more scarce, certainly when the goal of the experiment is to document the biomechanical characteristics of explants. For this reason, there was a move from rabbits to sheep8. Adult ewes are large-animal models with a reasonably sized and accessible vagina. They can be used for the mid-term evaluation of novel implants, and it is possible to reproduce vaginal exposures with certain materials9,10,11,12,13. Not only the dimensions and anatomy of the ovine vagina and pelvic floor are comparable to those in humans, but also the spontaneous occurrence of prolapse, which occurs in 15% of ewes. Prolapse risk factors are overlapping (i.e., multiparity, previous history of POP, increased intra-abdominal pressure induced by a higher bodyweight or when grazing on hills, and comparable effects of (phyto)estrogens)6,14. In Europe, sheep are the only reasonable alternative, as research on non-human primates has been nearly completely banned. Here, the model was taken one step further by mimicking the transvaginal insertion of implants using trocars and guides for the tension-free placement of meshes into the recto-vaginal septum. This was followed by fixing the implant using anchoring with arms through the ligaments of muscles, which can be considered equivalent to clinical practice15,16. So far, this technique has not been studied, though many believe that specific complications may occur due to the use of these longer strips and/or the piercing of anatomical structures.
In an earlier detailed anatomical study, the ovine pelvic floor was compared to the female pelvis17. When it comes to anchoring the implant, sheep do not have the sacrospinous ligament, yet they do have a very well-developed and broad sacrotuberous ligament. The pudendal nerve runs ventrally over it, making it unsafe to use this landmark as a suspension point. Conversely, the coccygeus muscle and its fascia, as well as the obturator membrane, are accessible through the rectovaginal space. Here, the access and position of the anatomical structures for the fixation of anchoring arms is proposed. The instruments that can be used to position the mesh are discussed. Finally, the relationship of the arms or trocars to adjacent anatomical structures, such as vessels and nerves, as well as potential intraoperative complications, are also described.

<table><tbody><tr><td>Animals:&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>parous female sheep (45 - 65 kg)</td><td>Zo&amp;ouml;technical Institute of the KU Leuven</td><td>NA</td><td>experimetnal animal</td></tr><tr><td>Sterile clothing:&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>sterile drape 45 x 75 cm</td><td>Lohmann &amp;amp; Rauscher, Regensdorf, Germany</td><td>33002</td><td>other material</td></tr><tr><td>sterile OR drape 150 x 180 cm</td><td>Lohmann &amp;amp; Rauscher, Regensdorf, Germany</td><td>33009</td><td>other material</td></tr><tr><td>sterile glowes 2x</td><td>Lohmann &amp;amp; Rauscher, Regensdorf, Germany</td><td>16652</td><td>other material</td></tr><tr><td>sterile surgical gown 2x</td><td>Lohmann &amp;amp; Rauscher, Regensdorf, Germany</td><td>19342</td><td>other material</td></tr><tr><td>surgical head cap 2x</td><td>Lohmann &amp;amp; Rauscher, Regensdorf, Germany</td><td>17427</td><td>other material</td></tr><tr><td>surgical face mask 2x</td><td>Lohmann &amp;amp; Rauscher, Regensdorf, Germany</td><td>11983</td><td>other material</td></tr><tr><td>Other surgical material</td><td></td><td></td><td></td></tr><tr><td>implant</td><td>FEG Textiltechnik GmbH, Aachen, Germany</td><td>NA</td><td>purposely designed implant</td></tr><tr><td>3/0 polypropylene suture</td><td>Prolene, Ethicon, Diegem, Belgium</td><td>8762H</td><td>suture material</td></tr><tr><td>3/0 polygecaprone suture</td><td>Vicryl, Ethicon</td><td>J311H</td><td>suture material</td></tr><tr><td>gauze swabs 10 x 10 cm 10x, 12-ply</td><td>Lohmann &amp;amp; Rauscher, Regensdorf, Germany</td><td>11574</td><td>other material</td></tr><tr><td>syringe 10 mL</td><td>Becton Dickinsosn S.A., Madrid, Spain</td><td>300613</td><td>aqua-dissection</td></tr><tr><td>needle 16 gauge</td><td>Terumo, Leuven, Belgium</td><td>NN-2238R</td><td>aqua-dissection</td></tr><tr><td>Surgical equipment:</td><td></td><td></td><td></td></tr><tr><td>blade no.22</td><td>Fine science isntruments, Heidelberg, Germany</td><td>10022-00</td><td>surgical instruments</td></tr><tr><td>Allis tissue forceps 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>11091-15</td><td>surgical instruments</td></tr><tr><td>Standart pattern forceps 1 x 2 theeth 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>11023-14</td><td>surgical instruments</td></tr><tr><td>Standart pattern forceps straight serrated 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>11000-14</td><td>surgical instruments</td></tr><tr><td>Scalpel handle 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>10004-13</td><td>surgical instruments</td></tr><tr><td>Halstead-Mosquito forceps 2x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>13008-12</td><td>surgical instruments</td></tr><tr><td>Standart pattern scissors 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>14001-14</td><td>surgical instruments</td></tr><tr><td>Metzenbaum scissors 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>14016-18</td><td>surgical instruments</td></tr><tr><td>Crile Wood needle holder 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>12003-15</td><td>surgical instruments</td></tr><tr><td>Kell forceps 1x</td><td>Fine science isntruments, Heidelberg, Germany</td><td>13018-14</td><td>surgical instruments</td></tr><tr><td>Long Starr Self-Retaining Retractor with eight 5 mm sharp stay hooks</td><td>Cooper Surgical, Tumbull, USA</td><td>3704</td><td>surgical instruments</td></tr><tr><td>Heaney Simon Vaginal Retractor</td><td>Medical supplies &amp;amp; equipments co., Katy, Texas, USA</td><td>403-129FSI</td><td>surgical instruments</td></tr><tr><td>Trocar (Insnare)</td><td>Bard, West Sussex, United Kingdom</td><td>NA</td><td>any trocar on market for transvaginal mesh implantation</td></tr><tr><td>Medication:</td><td></td><td></td><td></td></tr><tr><td>amoxilicilline clavulanate 1000 mg / 300 mL (Ampiciline)</td><td>GSK, Wavre, Belgium</td><td>NA</td><td>antibiotics</td></tr><tr><td>buprenorfin 0.3 mg/mL + chlorocresol 1.35 mg/mL (Vetregesic)</td><td>Ecuphar, Oostkamp, Belgium</td><td>NA</td><td>analgesia</td></tr><tr><td>ketamin HCL 100 mg/mL (Ketamine 1000)</td><td>Ceva Sante Animale, Brussels, Belgium</td><td>NA</td><td>anesthesia</td></tr><tr><td>isoflurane (IsoFlo)</td><td>Abbott Laboratories Ltd, Maidenhead, Berkshire, UK</td><td>NA</td><td>anesthesia</td></tr><tr><td>polyvidone iodium 7.5% (Braunol)</td><td>B. Braun Medical, Machelen, Belgium</td><td>NA</td><td>local desinfection</td></tr><tr><td>saline solution 500 mL</td><td>B. Braun Medical, Machelen, Belgium</td><td>NA</td><td>aqua-dissection</td></tr><tr><td>Xxylazine HCl , 1 mL/50 kg&amp;nbsp;</td><td>Vexylan, Ceva Sante Animale, Belgium</td><td>NA</td><td>premedication</td></tr><tr><td>atropine Sulfate 15 mg/mL (),</td><td>Viatris, Belgium</td><td>NA</td><td>premedication</td></tr></tbody></table>

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.

Management in a Longer Observation Setup
Following the surgical procedure, vaginal packing (a saline-solution-soaked gauze package inserted in the vagina immediately after the surgery) may be inserted for 24 h to secure the implant position. The sheep should be placed in a recovery cage and its respiratory function followed until full recovery. Later, it is possible to place the sheep in the stable and to allow it to move freely and to drink and eat ad libitum. The vaginal packing, if present, must be removed 24 h after the surgery. The sheep should receive analgesics (buprenorfin and chlorocresol, 1 mL, i.m.) for at least three postoperative days. During the first postoperative week, the animal should be checked daily, and then every week until the end of the experiment.
Surgical Feasibility
During the procedure, there were no problems with mesh insertion in any of the animals. There was almost no bleeding during the dissection of the rectovaginal septum and paravaginal spaces. It was possible to identify the medial aspect of the obturator foramen through the dissection. Also, the trocar insertion was straightforward, with a difference in resistance between the more compliant muscles and the more resistant fascia of the individual muscles. Though the initial trajectory through the muscles was less controlled, the tip of the trocar became palpable once in contact with the obturator muscle. The dorsal arms were placed without complications or obstacles.
Mesh Positioning and Findings during Subsequent Dissections
To investigate the proper positioning of the implant with the anchoring arms, which was considered more difficult to achieve, three animals were euthanized by an intravenous injection of 1.0 mL of an embutramide-mebezonium-tetracaine hydrochloride mixture. Following euthanasia, the surgical area was carefully dissected to explore the mesh position and the effect of the insertion of the arms of the mesh through the relevant anatomical structures. The shortest distance of the ventral arms from the obturator artery and nerve to the internal pudendal vessel was measured with a ruler, and their relationship to the tendinous arc of the levator ani was investigated. Dorsally, the pudendal nerve and internal pudendal artery were identified, and the distance to them measured with a ruler.
Relevant bleeding did not occur in any of the ewes, nor was there any intra-operative nerve, intestinal, or bladder injury. In the first sheep, the cranial arm passed through the caudal aspect of the cul-de-sac, but the bowels remained intact. This was avoided in the next sheep by guiding the tip of the trocar more laterally away from the cul-de-sac. The other arm passages were identified in the anatomical structures previously described, far away from the pelvic nerves and vessels. The cranial arms passed through the caudal aspect of the obturator foramen (Figure 4, panel A). The entry point of the trocar was at the level of the tendinous arc of the levator ani, 2 - 2.5 cm caudal to the obturator canal and the obturator vessels and nerve (Figure 4, panel B). Once in the paravaginal space, the arm was located 1 - 1.5 cm ventral to the pudendal artery and vein and 1 cm lateral to the vaginal artery. The caudal arms passed 1 cm caudal to the caudal aspect of the broad sacrotuberous ligament, right through the coccygeus muscle. In that location, there are no major vessels or nerves anywhere close by. The pudendal nerve is located on the inner surface of the caudal part of the sacrotuberous ligament.
The central part of the mesh was placed flat, with its cranial part stretching retroperitoneally under the caudal end of the cul-de-sac and its caudal part down along the rectovaginal septum. No rectal perforations occurred (Figure 4, panel D).
<img alt="Figure 4" src="/files/ftp_upload/55706/55706fig4.jpg" /> 
Figure 4. Anatomical Dissection on a Pelvic Hemisection. A: The lateral pelvic sidewall after the removal of the parietal peritoneum and the retroperitoneal fat tissue. The pubis (P) is at the top of the figure and the vagina (V) is moved medially to reveal the course of the cranial arm (full arrow). The arm passes through the tendinous arc of the levator ani. The green pin corresponds to the position of the internal pudendal vessels, the blue pin marks the vaginal artery, and the yellow pin is placed in the levator ani. B: The lateral pelvic sidewall after removal of the arm (course marked with a yellow line). The entrance point is located in the caudal aspect of the obturator foramen (marked with four blue pins). The obturator nerve and vessels (red line) pass through its cranial aspect. The internal pudendal and vaginal vessels (green and blue line, respectively) are dorsal to the arm. The obturator muscle (OM) and the levator ani muscle (LAM) are indicated as well. C: The course of the arm (open arrow) through the medial muscles of the thigh. The gracilis muscle is moved medially to display the course of the arm through the semitendinosus and the adductor magnus muscle. D: The central part of the implant (open arrow) is placed between the vagina (V), the parietal peritoneum, and the rectum (R).
<table fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td rowspan="2">Rectangular mesh</td>
			<td colspan="2">Arm mesh&#160;</td>
		</tr>
		<tr>
			<td></td>
			<td>Central body&#160;</td>
			<td>Arms</td>
		</tr>
		<tr>
			<td>Dimensions (mm x mm)</td>
			<td>30 x 40</td>
			<td>30 x 40</td>
			<td>10 x 150</td>
		</tr>
		<tr>
			<td>Thickness (mm)</td>
			<td>0.54</td>
			<td>0.54</td>
			<td>0.7</td>
		</tr>
		<tr>
			<td>Weight (g/m2)</td>
			<td>83</td>
			<td>83</td>
			<td>73</td>
		</tr>
		<tr>
			<td>Pore size (mm2)</td>
			<td>2.5 x 2.5</td>
			<td>2.5 x 2.5</td>
			<td>1.0 x 1.4</td>
		</tr>
		<tr>
			<td>Stiffness (N/mm)</td>
			<td>0.3</td>
			<td>0.3</td>
			<td>14.7</td>
		</tr>
		<tr>
			<td>Anisotropic index&#160;</td>
			<td>1.3</td>
			<td>1.3</td>
			<td>7.5</td>
		</tr>
	</tbody>
</table>
Table 1: Dry Material Properties. 
The table shows the material properties of the rectangular mesh and the mesh with anchoring arms. The stiffness and anisotropic index were obtained from Maurer et al18.

Watch this Scientific Journal Video about Transvaginal Mesh Insertion in the Ovine Model at JoVE.com

Transvaginal Mesh Insertion in the Ovine Model

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 ...

transvaginal-mesh-insertion-in-the-ovine-model

Universidad San Sebastian

Research

JoVE Journal

Medicine

15.9K Views.  Clinical Specialties Research Groups, Faculty of Medicine, KU Leuven. 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. 

Video: Transvaginal Mesh Insertion in the Ovine Model

An Orthotopic Endometrial Cancer Model with Retroperitoneal Lymphadenopathy Made From In Vivo Propagated and Cultured VX2 Cells

Endometrial cancer is the most common gynecologic malignancy in North America and the incidence is rising worldwide. Treatment consists of surgery with or without adjuvant therapy depending on lymph node involvement as determined by lymphadenectomy. Lymphadenectomy is a morbid procedure, which has not been shown to have a therapeutic benefit in many patients, and thus a new method to diagnose lymph node metastases is required. To test novel imaging agents, a reliable model of endometrial cancer with retroperitoneal lymph node metastases is needed. The VX2 endometrial cancer model has been described frequently in the literature; however, significant variation exists with respect to the method of model establishment. Furthermore, no studies have reported on the use of cultured VX2 cells to create this model as only cells propagated in vivo have been previously used. Herein, we present a standardized surgical method and post-operative monitoring method for the establishment of the VX2 endometrial cancer model and report on the first use of cultured VX2 cells to create this model.

This protocol presents a standardized method to grow VX2 cells in culture and to create an orthotopic VX2 model of endometrial cancer with retroperitoneal lymph node metastases in rabbits. Orthotopic endometrial cancer models are important for the pre-clinical study of novel imaging modalities for the diagnosis of lymph node metastases.

Endometrial cancer is the most common gynecologic malignancy in North America and the incidence is rising worldwide. Treatment consists of surgery with or ...

Cancer Research

Anterior Cervical Discectomy and Fusion in the Ovine Model

Anterior cervical discectomy and fusion (ACDF) is the most common surgical operation for cervical radiculopathy and/or myelopathy in patients who have failed conservative treatment1,5. Since the operation was first described by Cloward2 and Smith and Robinson6 in 1958, a variety refinements in technique, graft material and implants have been made3. In particular, there is a need for safe osteoinductive agents that could benefit selected patients. The ovine model has been shown to have anatomical, biomechanical, bone density and radiological properties that are similar to the human counterpart, the most similar level being C3/44. It is therefore an ideal model in which preclinical studies can be performed. In particular this methodology may be useful to researchers interested in evaluating different devices and biologics, including stem cells, for potential application in human spinal surgery.

This video demonstrates the technique of anterior cervical discectomy and fusion in the ovine model.

Anterior cervical discectomy and fusion (ACDF) is the most common surgical operation for cervical radiculopathy and/or myelopathy in patients who have ...

Biology

Methods for Studying Uterine Contributions to Pregnancy Establishment in an Ovariectomized Mouse Model

For pregnancy to be established, a viable blastocyst must successfully interact with a receptive uterine lining (endometrium) to facilitate implantation and placenta formation and enable ongoing pregnancy. The limitations to pregnancy success caused by embryonic defects are well known and have been largely overcome in recent decades with the rise of in vitro fertilization (IVF) and assisted reproductive technologies. As yet, however, the field has not overcome the limitations caused by an inadequately receptive endometrium, thus resulting in stagnating IVF success rates. Ovarian and endometrial functions are closely intertwined, as hormones produced by the ovary are responsible for the endometrium's menstrual cyclicity. As such, when using rodent models of pregnancy, it can be difficult to ascertain whether an observed result is due to an ovarian or uterine deficit. To overcome this, an ovariectomized mouse model was developed with embryo transfer or artificial decidualization to allow the study of uterine-specific contributions to pregnancy. This article will provide instructions on how to perform ovariectomy and offer insights into various techniques for supplying exogenous hormones to support successful artificial decidualization or pregnancy following embryo transfer from healthy donors. These techniques include subcutaneous injection, slow-release pellets, and osmotic mini pumps. The key advantages and disadvantages of each method will be discussed, enabling researchers to choose the best study design for their specific research question.

Pregnancy establishment is a dynamic process involving complex embryo and uterine crosstalk. The precise contributions of the maternal uterine environment to these processes remain an active area of investigation. Here, detailed protocols are provided to aid in designing in vivo animal models to address these research questions.

For pregnancy to be established, a viable blastocyst must successfully interact with a receptive uterine lining (endometrium) to facilitate implantation ...

Developmental Biology

In Vivo Real-Time Study of Drug Effects on Carotid Blood Flow in the Ovine Fetus

The ability of an organism to maintain a constant blood flow to the brain in response to sudden surges in systemic blood pressure (BP) is known as cerebral autoregulation (CAR), which occurs in the carotid artery. In contrast to full-term neonates, preterm neonates are unable to reduce the cerebral blood flow (CBF) in response to increased systemic BP. In preterm neonates, this exposes the fragile cerebral vessels to high perfusion pressures, leading to their rupture and brain damage. Ex vivo studies using wire myography have demonstrated that carotid arteries from near-term fetuses constrict in response to the activation of adrenergic alpha1 receptors. This response is blunted in the preterm fetus. Thus, to examine the role of alpha1-AR in vivo, presented here is an innovative approach to determine the effects of drugs on a carotid arterial segment in vivo in an ovine fetus during the developmental progression of gestation. The presented data demonstrate the simultaneous measurement of fetal blood flow and blood pressure. The perivascular delivery system can be used to conduct a long-term study over several days. Additional applications for this method could include viral delivery systems to alter the expression of genes in a segment of the carotid artery. These methods could be applied to other blood vessels in the growing organism in utero as well as in adult organisms.

In Vivo Real-Time Study of Drug Effects on Carotid Blood Flow in the Ovine Fetus

The present protocol describes a method to deliver drugs and gene expression-modifying agents perivascularly in an in utero developing fetus. Importantly, the effect of drugs/agents on blood flow can be measured with the progression of pregnancy.

The ability of an organism to maintain a constant blood flow to the brain in response to sudden surges in systemic blood pressure (BP) is known as ...

Ovine Lumbar Intervertebral Disc Degeneration Model Utilizing a Lateral Retroperitoneal Drill Bit Injury

Intervertebral disc degeneration is a significant contributor to the development of back pain and the leading cause of disability worldwide. Numerous animal models of intervertebral disc degeneration have been developed. The ideal animal model should closely mimic the human intervertebral disc with regard to morphology, biomechanical properties and the absence of notochordal cells. The sheep lumbar intervertebral disc model fulfils these criteria. We present an ovine model of intervertebral disc degeneration utilizing a drill bit injury through a lateral retroperitoneal approach. The lateral approach significantly reduces the incision and potential morbidity associated with the traditional anterior approach to the ovine spine. Utilization of a drill-bit method of injury affords the ability to produce a consistent and reproducible injury, of precise dimensions, that initiates a consistent degree of intervertebral disc degeneration. The focal nature of the annular and nucleus pulposus defect more closely mimics the clinical condition of focal intervertebral disc herniation. Sheep recover rapidly following this procedure and are typically mobile and eating within the hour. Intervertebral disc degeneration ensues and sheep undergo necropsy and subsequent analysis at periods from eight weeks. We believe that the drill bit injury model of intervertebral disc degeneration offers advantages over more conventional annular injury models.

Intervertebral disc degeneration is a significant contributor to back pain and a leading cause of disability worldwide. Numerous animal models of intervertebral disc degeneration exist. We demonstrate an ovine model of intervertebral disc degeneration, utilizing a drill bit, which achieves a consistent disc injury and reproducible level of disc degeneration.

Intervertebral disc degeneration is a significant contributor to the development of back pain and the leading cause of disability worldwide. Numerous ...

Intravitreal Injections in the Ovine Eye

There are several methods for the delivery of therapeutic agents to the retina, including intravitreal (IVT), subretinal, suprachoroidal, periocular, or topical administration. IVT drug delivery involves an injection into the vitreous humor of the eye, a gelatinous substance that fills the posterior chamber of the eye and maintains the shape of the eye globe. Although the IVT route is less specifically targeted than subretinal delivery, it is much less invasive and is widely used in clinical settings for a range of ocular diseases. 
We previously demonstrated the efficacy of intravitreal delivery of an adeno-associated virus (AAV)-mediated gene therapy product (AAV9.CLN5) in sheep with a naturally occurring CLN5 form of neuronal ceroid lipofuscinosis (NCL). Affected sheep received IVT gene therapy in one eye, with the other untreated eye serving as an internal control. Retinal structure and function were maintained in the treated eye up to 15 months after treatment, while the untreated eye displayed progressively declining function and severe atrophy during postmortem examination. Based on the sheep studies, the CLN5 gene therapy product was cleared as a candidate investigational new drug (IND) by the United States Food and Drug Administration in September 2021. This paper details the surgical protocol for IVT delivery of a therapeutic viral vector to the ovine eye.

Intravitreal injections were performed in the sheep eye with the aim of delivering viral-mediated gene therapy to the retina.

There are several methods for the delivery of therapeutic agents to the retina, including intravitreal (IVT), subretinal, suprachoroidal, periocular, or ...

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 ...

Chronic Ovine Model of Right Ventricular Failure and Functional Tricuspid Regurgitation

The pathophysiology of severe functional tricuspid regurgitation (FTR) associated with right ventricular dysfunction is poorly understood, leading to suboptimal clinical results. We set out to establish a chronic ovine model of FTR and right heart failure to investigate the mechanisms of FTR. Twenty adult male sheep (6-12 months old, 62 &#177; 7 kg) underwent a left thoracotomy and baseline echocardiography. A pulmonary artery band (PAB) was placed and cinched around the main pulmonary artery (PA) to at least double the systolic pulmonary artery pressure (SPAP), inducing right ventricular (RV) pressure overload and signs of RV dilatation. PAB acutely increased the SPAP from 21 &#177; 2 mmHg to 62 &#177; 2 mmHg. The animals were followed for 8 weeks, symptoms of heart failure were treated with diuretics, and surveillance echocardiography was used to assess for pleural and abdominal fluid collection. Three animals died during the follow-up period due to stroke, hemorrhage, and acute heart failure. After 2 months, a median sternotomy and epicardial echocardiography were performed. Of the surviving 17 animals, 3 developed mild tricuspid regurgitation, 3 developed moderate tricuspid regurgitation, and 11 developed severe tricuspid regurgitation. Eight weeks of pulmonary artery banding resulted in a stable chronic ovine model of right ventricular dysfunction and significant FTR. This large animal platform can be used to further investigate the structural and molecular basis of RV failure and functional tricuspid regurgitation.

Right ventricular failure and functional tricuspid regurgitation are associated with left-sided heart disease and pulmonary hypertension, which contribute significantly to morbidity and mortality in patients. Establishing a chronic ovine model to study right ventricular failure and functional tricuspid regurgitation will help in understanding their mechanisms, progression, and possible treatments.

The pathophysiology of severe functional tricuspid regurgitation (FTR) associated with right ventricular dysfunction is poorly understood, leading to ...

Optimizing Oviductal Embryo Transfer in Cloned Pigs

Embryo Transfer Surgery via Laparotomy in Gilts

This protocol aims to demonstrate the surgical technique for transferring cloned pig embryos to the oviduct, a method widely used in the production of genetically modified pigs for biomedical research. Nine gilts underwent hormonal synchronization and laparotomy for the transfer of cloned embryos produced by somatic cell nuclear transfer (SCNT) at stages of up to 4 cells on day 2 to the oviduct. Gestational diagnosis was conducted via ultrasound examination 30 days post-transfer surgery. Six out of the nine operated gilts exhibited signs of pregnancy on ultrasound examination. However, as there was no progression in fetal development as assessed by echography, the gilts underwent necropsy at 60 days for the collection of biological material and assessment of the reproductive system. Adhesions were observed in the uterine horns, ovaries, and oviducts. From the uterine lumen of two of the euthanized gilts, one and four embryonic structures with gestational ages ranging between 12 to 20 days were obtained. Despite the absence of live piglets, likely attributed to the low-efficiency rate of transferring cloned pig embryos, which is influenced by various factors, including the number and quality of transferred embryos, the presented surgical technique proved to be rapid and safe.

Author Spotlight: Surgical Methods and Outcomes in Oviductal Cloned Pig Embryo Transfers

This protocol describes the application of the surgical technique used to transfer cloned pig embryos via laparotomy in gilts.

This protocol aims to demonstrate the surgical technique for transferring cloned pig embryos to the oviduct, a method widely used in the production of ...

Modeling Transuterine Fetal Tracheal Occlusion in Murine Model: A Surgical Procedure for Ligation of the Fetal Trachea Within a Pregnant Mouse

Source: Ayd&#305;n, E. et al. <a href="https://www-jove-com.bdigitaluss.remotexs.co/v/61772">Transuterine Fetal Tracheal Occlusion Model in Mice.</a> J. Vis. Exp. (2021)
This video demonstrates the transuterine tracheal occlusion procedure of a mouse fetus in a pregnant mouse. This model can be used to study the impact of occluded trachea on lung development.

Modelowanie przezmacicznej niedrożności tchawicy płodu w modelu mysim: zabieg chirurgiczny podwiązania tchawicy płodu u ciężarnej myszy

Source: Ayd&#305;n, E. et al. Transuterine Fetal Tracheal Occlusion Model in Mice. J. Vis. Exp. (2021)
This video demonstrates the transuterine tracheal ...

Transvaginal Mesh Insertion in the Ovine Model

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Embryo Transfer Surgery via Laparotomy in Gilts

Modelowanie przezmacicznej niedrożności tchawicy płodu w modelu mysim: zabieg chirurgiczny podwiązania tchawicy płodu u ciężarnej myszy