The authors would like to acknowledge the Administration of the FPO-IRCCS of Candiolo (Turin) for the contribution and the constant support to our work.

1. Collection of the Specimens
<ol>
	<li>Take lamb and porcine innards from animals slaughtered for meat products. 
	NOTE: Innards should be supplied by a reliable butcher who has complied with current health standards.</li>
	<li>Collect the larynx together with the base of the tongue and the first five tracheal rings, in order to give stability to the specimen. Leave the remaining innards with the butcher, especially the brain and spinal cord, to avoid infective tissue.</li>
	<li>Thoroughly wash the specimen and put it in a numbered box for tracking.</li>
	<li>Use the specimen right away or freeze it at -18 &#176;C and defrost it at least 24 h before the dissection.</li>
</ol>
2. Preparation of the Laboratory
<ol>
	<li>If possible, use a sector table with a proper sink, easily washable before and after use.</li>
	<li>Procure a surgical light or a traditional lamp providing sufficient illumination.</li>
	<li>Put a barrier across the table at the halfway point to split it into two stations. 
	NOTE: This allows for more apprentices to work at the same time and protects them from the laser beam.</li>
	<li>Procure a special waste container where the specimen and used parts will eventually be discarded. Close the container, label it with the specific European Waste Catalogue (EWC) code, and dispose of it according to the institution&#39;s protocols.</li>
	<li>Optionally, set the air conditioning to offset heat from all of the machinery working and maintain a steady temperature in the room.</li>
</ol>
3. Preparation of the Endoscopic Station
<ol>
	<li>Put the specimen on a proper support, positioning the laryngoscope at the end of the surgical table (Figure 1). 
	NOTE: The stand support is the one proposed by Delfo Casolino and Andrea Ricci Maccarini and is made of a metal folding structure with adjustable transverse bars23. The stand support is equipped with a holder for a laryngoscope and a chassis for specimen positioning.</li>
	<li>For the safety of the attendants, position an open wood box around the station in order to absorb potential improperly directed laser beams. 
	NOTE: Recently the literature has reported a new validated station, approved for CO2 laser surgery, called LarynxBox. This transparent structure is entirely made by polycarbonate resin, which can safely absorb all the misdirected laser beams14,24 and could be a replacement for the wooden structure described above.</li>
	<li>Insert the laryngoscope inside the specimen, expose the desired surgical target (i.e., supraglottic, arytenoid, glottic plane, etc.) and fix the laryngoscope to the support by tightening the proper screw. 
	NOTE: Be sure for safety and for the precision of the surgery to firmly fix both the laryngeal specimen and the laryngoscope to the metal structure. If not, use dedicated needles or tape to steadily secure the specimen to the station.</li>
	<li>Choose the proper laryngoscope for the selected laryngeal region. For example, use a wide and curved laryngoscope for the supraglottic region (i.e., Lindholm operating laryngoscope), a straight and narrow one for the vocal folds (i.e., Dedo operating laryngoscope).</li>
	<li>For exposure of the vocal folds in the porcine specimen, embed the tip of the laryngoscope anterior to the arytenoid cartilages, pushing these structures in a lateroposterior direction, in order to open and put tension on the vocal folds.</li>
	<li>Place a suction system inside the specimen, from above or below, in order to extract the laser fumes.</li>
	<li>Put and fix in position a wet gauze inside the inferior tracheal extremity of the larynx, in order to avoid the emission of CO2 laser from the inferior part of the specimen. In the same way, place a wet gauze at the superior border of the larynx in order to protect the areas not involved in the dissection.</li>
	<li>Connect the operating microscope to the CO2 laser, and place it on the right side of the table.</li>
	<li>Ensure that the surgeon and all the participants wear safety goggles before turning on the CO2 laser.</li>
	<li>Put the endoscope or the exoscope in front of the laryngeal specimen in order to guarantee that the attendants get the same perspective as that of the first operator. 
	NOTE: Make sure to put the endoscope or the exoscope above the CO2 laser source in order to avoid a collision between the laser and the instrumentation.</li>
	<li>Use an endoscope holder to maintain the optic system in place. Ensure that all the components are fixed and stable in their position for safety and for the surgical dissection.</li>
	<li>Put the 4k or full high definition (FHD) monitor on the left side of the table, linked to the microscope or to the endoscopic camera.</li>
	<li>Prepare a microlaryngoscopy set of surgical instruments on a table, beside the first surgeon. 
	NOTE: The set should at least contain laryngeal forceps, scissors and spreader, telescope with light cable, ball end suction device, laryngeal hook, laryngeal needle, gauzes.</li>
	<li>Start the dissection.</li>
</ol>
4. Preparation of the Open Surgery Station
<ol>
	<li>Place the specimen on the other end of the sector table, inside an open box.</li>
	<li>Prepare the open laryngeal surgery set on a table beside the operating field. 
	NOTE: The set should at least be composed of scissors, a pair of forceps (traumatic and atraumatic), dissector, scalpels, pin cutter, hook, needle holder, and stitches.</li>
	<li>Adjust the surgical light so it is on the surgical field.</li>
	<li>Optionally, set the CO2 fiber laser device.</li>
	<li>Optionally, put a conventional two-dimensional (2D) camera or a three-dimensional (3D) exoscope above the surgical field and connect it to a 2D/3D monitor. 
	NOTE: The tutor and the other surgeons can watch what the operator is doing and provide guidance while wearing polarized glasses.</li>
</ol>
5. (Optional) Broadcast the Dissection
<ol>
	<li>Set up an ambient camera, capable of filming all of the room.</li>
	<li>Link the two monitors, used in the dissections, to a workstation.</li>
	<li>Broadcast the signal to an external room to extend the procedure to the public, to make comments, or guide the dissection remotely.</li>
</ol>
6. Endoscopic Dissection
<ol>
	<li>Start with a bilateral vestibulectomy in order to improve the view on the glottic plane. Turn on the CO2 laser and use a 6&#8722;10 W power, SuperPulse or UltraPulse mode, a length of 0.8&#8722;1.5 mm and a depth of 1&#8722;2 points. Use the micromanipulator in order to move the laser pointer and micro forceps in order to grab the mucosa while performing the vestibulectomy.</li>
	<li>Once the vestibulectomy is performed, inject 2 mL of a NaCl solution (0.9%) into Reinke&#39;s space in order to highlight the mucosa.</li>
	<li>Perform the superior cordotomy: using the CO2 laser or the microscissors incise the mucosa longitudinally along the superior and lateral aspect of the vocal cord. Grasp the mucosa with forceps and dissect Reinke&#39;s space in order to identify the underlying vocalis muscle.</li>
	<li>Perform a bilateral cordectomy, from type I to V, based on the goal of the dissection, according to the European Laryngological Society (ELS) classification by Remarcle et al.25. 
	NOTE: If a porcine model is used for endoscopic dissection, it is not possible to perform a type II cordectomy, because the vocal ligament is absent. Cordectomies can be performed both by CO2 laser (4&#8722;6 W, Super or Ultrapulse mode, a length of 0.8&#8722;1.5 mm, and a depth of 1&#8722;2 points) or by cold surgical instrumentations (endoscopic microforceps and microscissors).</li>
	<li>Once the cordectomy is performed, extract the surgical specimen and place it on a working table. Try to define the anatomical landmarks (e.g., anterior, posterior, and deep aspects).</li>
	<li>Approach the paraglottic spaces and dissect the region for any anatomical purpose, paying attention to the anatomical landmarks and limits.</li>
	<li>Perform supraglottic laryngectomies from I to IVb according to the classification of Remacle et al.26 and approach the pre-epiglottic space. 
	NOTE: It must be kept in mind that porcine larynges have larger arytenoids and a smaller epiglottis than in humans.</li>
</ol>
7. Open Dissection I (OPHL)
<ol>
	<li>Dissect the strap muscles along the midline using scissors and forceps.</li>
	<li>Remove the prelaryngeal tissue.</li>
	<li>For larynx skeletonization, rotate the larynx contralaterally and perform, using scissors or a scalpel, an incision of the inferior constrictor muscle bilaterally along the lateral aspect of the thyroid cartilage. This procedure can also be performed with a CO2 fiber laser27, if available. Protect the superior laryngeal pedicle by retracting the larynx medially and downwards, then section the thyrohyoid ligament. Bilaterally dissect the pyriform sinus from the thyroid cartilage and the paraglottic space, as far as the inferior cornu of the thyroid cartilage.</li>
	<li>Dissect the cricothyroid muscle and section the inferior cornu of the thyroid cartilage bilaterally to protect the recurrent laryngeal nerve.</li>
	<li>Manually fracture the thyroid cartilage along the midline. Push with the thumbs on the laryngeal prominence while pulling forward the lateral laminae of the cartilage.</li>
	<li>Using a scalpel, make the superior access along a line parallel to the superior border of the thyroid cartilage through the pre-epiglottic space. Remove the pre-epiglottic space. Modify the superior access according to the selected type of OPHL, following the ELS classification6.</li>
	<li>Using a scalpel, make the inferior access between the cricoid ring and the first tracheal ring. Modify the inferior access according to the selected type of OPHL, following the ELS classification6.</li>
	<li>Complete the dissection: using scissors or scalpel perform bilaterally the vertical incisions in order to connect the superior and the inferior accesses. Cut the ary-epiglottic folds, the false vocal cords, the true vocal cords, and the subglottic region. Modify the lines of incision according to the selected type of OPHL, following the ELS classification6.</li>
	<li>Perform the pexy: apply four polyglactin 910 stitches for OPHL types I and II, and six for OPHL type III, of which one median double, between the cricoid cartilage and the hyoid bone, passing through the base of the tongue. Ensure to make the passage of the lateral stitches adherent to the superior aspect of the hyoid bone in order to not damage the lingual artery. 
	NOTE: The inferior structure will vary based on the type of OPHL performed (thyroid cartilage for OPHL type I, cricoid cartilage for OPHL type II, first tracheal ring for OPHL type III).</li>
	<li>Optionally, check the result in the inside-out technique using a 0&#176; endoscopic telescope.</li>
</ol>
8. Open Dissection II (Total Laryngectomy)
<ol>
	<li>Remove the infrahyoid muscles using the scissors.</li>
	<li>Divide the thyroid gland isthmus and move the lobes off the trachea, cricoid, and inferior constrictor muscles.</li>
	<li>Rotate the larynx contralaterally and incise the inferior constrictor muscle along the lateral aspect of the thyroid cartilage using scissors or scalpel. Expose the pyriform sinus bilaterally. Release the greater cornu of the thyroid cartilage on both sides.</li>
	<li>Dissect bilaterally the pyriform sinus from the thyroid cartilage and the paraglottic space.</li>
	<li>Dissect the suprahyoid muscles from the hyoid bone following the superior border of the bone. 
	NOTE: Because in human patients the hypoglossal nerve and lingual artery are located at a depth below the greater cornu of the hyoid bone, simulate the maneuver by cutting the muscle insertion close to the medial aspect of the cornu.</li>
	<li>Perform the pharyngotomy through the valleculae, the pyriform sinus, or the postcricoid area. The choice of entry point is based on the size of the tumor. Use the scissors or the scalpel for this step.</li>
	<li>For inferior access, use the scalpel to incise the trachea between two tracheal rings and extend the tracheal incision posterolaterally.</li>
	<li>To perform the laryngectomy in the craniocaudal direction, begin from the epiglottis and proceed through the pharyngotomy. Using the scissors, cut the aryepiglottic folds then proceed through the lateral wall of the pyriform sinus. Incise the postcricoid mucosa transversely, dissecting the plane between the trachea and the esophagus. Remove the larynx.</li>
	<li>To perform the laryngectomy in a retrograde manner, use the scissors to transect the posterior membranous tracheal wall, dissecting above the trachea from the anterior esophageal wall. Incise the hypopharyngeal mucosa below the upper border of the cricoid lamina. Extend the incision to the pyriform sinus and remove the larynx.</li>
	<li>Perform the primary closure of the pharynx using interrupted absorbable sutures or barbed sutures in the horizontal direction. 
	NOTE: The sutures should be located submucosally on the external surface in order to avoid granulation and possible fistulas. The primary closure of the defect can be achieved easily if at least 2 cm of pharyngeal mucosa is preserved, otherwise a flap has to be harvested.</li>
</ol>

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The authors have nothing to disclose.

This paper aims to describe the organization of a laboratory dedicated to laryngeal surgery and the choice of equivalent ex vivo animal models that can be used to simulate several surgical procedures in an economical but faithful manner. When human specimens are not available, it is necessary to find an accurate animal model to be used as a substitute. If there is no anatomy department that can provide specimens from body donations, the average price for a human model is about $1,300&#8211;1,500. On the other hand, for an animal slaughtered for meat products, the equivalent ex vivo animal models are about $8 or less. Here, the experiences of setting up the dedicated space, individual training sessions, and the organization of surgical dissection courses are reported. Based on the literature, it was decided to use porcine and ovine laryngeal models, mainly for laser and open surgery, respectively10,14,15,19,20,21. Both the animal models described are easily available and affordable since they are animal waste products in the meat supply chain. Moreover, these ex vivo models are easily managed and stored, with no risk for the operators. Even if slightly different from the human larynx and removed from the normal context of the neck, the anatomical proportions and tissue composition of the animal substitutes are very similar, allowing a step-by-step reproduction of TLM, OPHL, and TL techniques. The large number of specimens available for a very reasonable price guarantees the possibility to repeat the procedure many times. In this way, surgeons can not only improve their precision and accuracy in surgical procedures, but they can also increase their speed of execution, principally during the less important surgical steps of the procedures.
The contemporary use of microscopes/endoscopes for the endolaryngeal view, together with the external view, enhanced in this case by the 3D exoscope, allows an inside-out perspective to be gained, which can help surgeons to fully understand the complex laryngeal anatomy and the importance of each surgical step. Moreover, the use of a camera and screen to share the dissection allows the tutor and the other surgeons to monitor the same field of view as the first operator, increasing the training potential of the system. In this way the tutor can guide the procedure, correct mistakes, and answer any questions or comments.
This type of set-up can be easily replicated, as it is modular and flexible based on the instruments and devices available. Naturally, possible limitations of the animal models can be found in the intrinsic differences between the model and the human larynx and in working on a single prepared organ in the absence of the normal relationships with the surrounding anatomical structures. In detail, the porcine larynx has different arytenoids conformation, which requires a good glottic exposure. Moreover, the absence of the vocal ligament in the porcine specimen prevents a completely realistic type II cordectomy. On the other hand, these differences are somewhat overshadowed by the availability and cost of the animal models, which are very similar substitutes in tissue consistency and structure. Once the surgeon has acquired sufficient ability, the natural step forward is to switch to simulation to the more expensive human specimens.
A laryngeal training center with the features described is an ideal set-up for training in this precision surgery, for technical refinement, and for teaching purposes. Moreover, the same laboratory can be used to test novel head and neck surgical techniques. For example, the growing diffusion of transoral robotic surgery for oropharyngeal and supraglottic tumors requires time for individual training on the robotic console and to experience tissue manipulation and movements. All of these exercises can be easily simulated and repeated inexpensively in a training laboratory organized as described, without moving surgical facilities and instruments.

In recent years, the field of laryngeal oncology has seen the introduction and spread of organ sparing protocols such as chemoradiotherapy (CRT), function sparing procedures like transoral laser microsurgery (TLM) and partial laryngectomies, and mainly open partial horizontal laryngectomies (OPHLs). Due to the current general propensity to give greater priority to a patient&#39;s quality of life after treatment, this strategy change was necessary in order to avoid, when possible, the burdensome consequences of the total laryngectomy (TL) procedure, which still remains the standard treatment for locally advanced laryngeal cancer. However, despite surgical and technical innovations, TL remains the ideal treatment for advanced stage laryngeal cancer (LC) and for patients who cannot tolerate a conservative protocol because of age or important comorbidities. Therefore, TL has to be properly included in the armamentarium of a complete laryngeal surgeon.
A relevant problem in learning about LC treatment is the relatively rare incidence of the pathology (~13,000 new diagnoses per year in the USA), against the wide spectrum of possible alternatives1,2. Moreover, as clearly stressed by Olsen in one of his editorials, the misinterpretation of studies that satisfy the standard of care leads to several unintended consequences3. One such consequence was the abandonment of TLM and OPHLs, because they were not included in those studies and in the cost-benefit evaluation, and therefore are no longer taught to residents and young surgeons3. As a result, there is a significant paucity of centers in which it is possible to actively learn a surgical technique demanding a high level of accuracy, where the difference between a conservative and an extirpative procedure is quantifiable in the order of millimeters.
In response to this background and to meet the need for dissemination of these surgical procedures, the European Laryngological Society has worked to standardize and classify both TLM and OPHL techniques4,5,6. The tremendous result of these classifications was to introduce the possibility of a modular treatment for LC, customized by the real tumor extent and always remaining inside the field of &#39;partial&#39; surgery and function sparing treatment.
As emphasized in recent work, surgical ability (as a matter of fact, the success of a procedure requires millimetric accuracy) and strict patient selection are mandatory for good results7,8,9. In good hands, and if applied to the proper patients and diseases, TLM and OPHL exhibit solid surgical and survival outcomes.
The practice and evolution of these surgical procedures took place almost exclusively in referral centers for pathology, due to the relatively high number of patients, which allowed the surgeons to develop the essential expertise to successfully treat even locally advanced LCs. Trying to summarize the current scenario, laryngeal surgery can be applied to a relatively small number of patients and consists of different procedures that are not available and viable in every center. To preserve laryngeal function and equally reach the oncological radicality, perfect comprehension of the geometrical anatomy, technical accuracy, and concern for the tissues, are mandatory. For all these reasons, simulations on models are nowadays necessary to successfully approach this type of surgery. Faithful, detailed simulations are required to consolidate the comprehension of the laryngeal framework, manage tissue manipulation with different techniques, and to learn the exact and precise sequence of movements required by a single procedure. Therefore, to learn TLM and OPHL techniques, it is appropriate to be able to practice in a dedicated laboratory. Where there is no possibility to train on human specimens, for ethical, economic, or availability reasons, it is necessary to find an alternative and affordable ex vivo model. Porcine and ovine larynges, waste animal products in the meat supply chain, are ideal and affordable models to simulate laryngeal surgery given their similarity to the human larynx in anatomical layout and tissue composition10,11.
Several groups have reported their experiences with porcine larynx used as a model for TLM11,12,13,14. Despite the different dimensions of the cartilaginous skeleton with larger arytenoids and the inability to distinguish between arytenoid, corniculate, and cuneiform cartilage, the glottic plane is very similar to its human counterpart: the arytenoid cartilage has an analogous articulation with the cricoid and similar geometric proportions15. When compared to other animal species, the porcine larynx has a defined laryngeal ventricle with well represented false vocal cords, while the glottic plane is characterized by short arytenoid vocal processes, long vocal folds, and the absence of a proper vocal ligament14. Furthermore, from the histologic point of view, Hahn and colleagues have reported a comparable elastin distribution within the lamina propria between porcine and human glottic planes16,17,18.
On the other hand, other studies have described the utilization of lamb larynx for both TLM and open surgeries10,19,20. In detail, Nisa et al. confirmed the strong similarity between ovine and human larynges, with the exception of a differently shaped hyoid bone and arytenoid cartilage, a lower position of the anterior commissure (placed at the inferior border of the thyroid cartilage), and near-complete tracheal rings21. Despite these small differences, those authors outlined the great utility of this model for training and practice of laryngotracheal surgical procedures21. Moreover, the same model was also used to simulate the percutaneous tracheostomy procedure22.
The aim of the present study is to illustrate how to prepare and organize a reproducible laboratory for laryngeal surgery on affordable and closely similar ex vivo animal laryngeal models. The authors&#39; experience in setting up such a laboratory was acquired during years of training on surgical simulation in a laboratory of experimental laryngeal surgery called &#34;Lary-Gym&#34; &#8211; at the FPO-IRCCS Cancer Institute of Candiolo, Turin, Italy.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3D camera</td>
			<td>STORZ</td>
			<td>VITOM 3D TH200</td>
			<td></td>
		</tr>
		<tr>
			<td>4k camera</td>
			<td>STORZ</td>
			<td>TH120</td>
			<td></td>
		</tr>
		<tr>
			<td>4K/3D 32&#34; monitor</td>
			<td>STORZ</td>
			<td>TM350</td>
			<td></td>
		</tr>
		<tr>
			<td>Autostatic arm for VITOM 3D</td>
			<td>STORZ</td>
			<td>28272 HSP</td>
			<td></td>
		</tr>
		<tr>
			<td>Bone Rongeur, Luer</td>
			<td>MEDICON</td>
			<td>30.30.35</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 fiber laser</td>
			<td>LUMENIS</td>
			<td>Ultrapulse/Surgitouch</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 laser</td>
			<td>LUMENIS</td>
			<td>AcuPulse 40WG</td>
			<td></td>
		</tr>
		<tr>
			<td>Dedo operating larygoscope</td>
			<td>STORZ</td>
			<td>8890 A</td>
			<td></td>
		</tr>
		<tr>
			<td>Delicate tissue forceps, Adson</td>
			<td>MEDICON</td>
			<td>06.21.12</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemostatic forceps curved</td>
			<td>MEDICON</td>
			<td>15.45.12</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemostatic forceps straight</td>
			<td>MEDICON</td>
			<td>15.44.12</td>
			<td></td>
		</tr>
		<tr>
			<td>Hook</td>
			<td>MEDICON</td>
			<td>20.48.05</td>
			<td></td>
		</tr>
		<tr>
			<td>Hopkins II forward-oblique telescope 30&#176;</td>
			<td>STORZ</td>
			<td>8712 BA</td>
			<td></td>
		</tr>
		<tr>
			<td>Hopkins II forward-oblique telescope 70&#176;</td>
			<td>STORZ</td>
			<td>8712 CA</td>
			<td></td>
		</tr>
		<tr>
			<td>Hopkins II straight forward telescope 0&#176;</td>
			<td>STORZ</td>
			<td>8712 AA</td>
			<td></td>
		</tr>
		<tr>
			<td>Image 1 pilot</td>
			<td>STORZ</td>
			<td>TC014</td>
			<td></td>
		</tr>
		<tr>
			<td>Kleinsasser handle</td>
			<td>STORZ</td>
			<td>8597</td>
			<td></td>
		</tr>
		<tr>
			<td>Kleinsasser hook 90&#176;</td>
			<td>STORZ</td>
			<td>8596 C</td>
			<td></td>
		</tr>
		<tr>
			<td>Kleinsasser injection needle straight</td>
			<td>STORZ</td>
			<td>8598 B</td>
			<td></td>
		</tr>
		<tr>
			<td>Kleinsasser scissors curved to left</td>
			<td>STORZ</td>
			<td>8594 D</td>
			<td></td>
		</tr>
		<tr>
			<td>Kleinsasser scissors curved to right</td>
			<td>STORZ</td>
			<td>8594 C</td>
			<td></td>
		</tr>
		<tr>
			<td>Kleinsasser scissors straight</td>
			<td>STORZ</td>
			<td>8594 A</td>
			<td></td>
		</tr>
		<tr>
			<td>Light source</td>
			<td>STORZ</td>
			<td>TL300</td>
			<td></td>
		</tr>
		<tr>
			<td>Lindholm distending forceps</td>
			<td>STORZ</td>
			<td>8654 B</td>
			<td></td>
		</tr>
		<tr>
			<td>Lindholm operating laryngoscope</td>
			<td>STORZ</td>
			<td>8587 A</td>
			<td></td>
		</tr>
		<tr>
			<td>Mayo standard scissors</td>
			<td>MEDICON</td>
			<td>03.50.14</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>LEICA</td>
			<td>F40</td>
			<td></td>
		</tr>
		<tr>
			<td>Module for 3D image</td>
			<td>STORZ</td>
			<td>Image 1 D3-link TC302</td>
			<td></td>
		</tr>
		<tr>
			<td>Module for 4K image</td>
			<td>STORZ</td>
			<td>Image 1 s 4U-Link TC304</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle Holder</td>
			<td>MEDICON</td>
			<td>10.18.65</td>
			<td></td>
		</tr>
		<tr>
			<td>Operating scissors standard curved</td>
			<td>MEDICON</td>
			<td>03.03.13</td>
			<td></td>
		</tr>
		<tr>
			<td>Raspatory, Freer</td>
			<td>MEDICON</td>
			<td>26.35.02</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractor, double-ended, Roux</td>
			<td>MEDICON</td>
			<td>22.16.13</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractor, Volkmann</td>
			<td>MEDICON</td>
			<td>22.34.03</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractory, double-ended, langenbeck</td>
			<td>MEDICON</td>
			<td>22.18.21</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel #11</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel #15</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Steiner Coagulation suction tube</td>
			<td>STORZ</td>
			<td>8606 D</td>
			<td></td>
		</tr>
		<tr>
			<td>Steiner Grasping forceps curved to left</td>
			<td>STORZ</td>
			<td>8663 CH</td>
			<td></td>
		</tr>
		<tr>
			<td>Steiner Grasping forceps curved to right</td>
			<td>STORZ</td>
			<td>8663 BH</td>
			<td></td>
		</tr>
		<tr>
			<td>Steiner Laryngoforce II grasping forceps</td>
			<td>STORZ</td>
			<td>8662 E</td>
			<td></td>
		</tr>
		<tr>
			<td>Steiner operating laryngoscope</td>
			<td>STORZ</td>
			<td>8661 CN</td>
			<td></td>
		</tr>
		<tr>
			<td>Suction tube to remove vapor</td>
			<td>STORZ</td>
			<td>8574 LN</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue grasping forceps</td>
			<td>MEDICON</td>
			<td>07.01.10</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Grasping forceps, Allis</td>
			<td>MEDICON</td>
			<td>50.02.15</td>
			<td></td>
		</tr>
		<tr>
			<td>Towel clamp</td>
			<td>MEDICON</td>
			<td>17.55.13</td>
			<td></td>
		</tr>
		<tr>
			<td>Vascular forceps, DeBakey</td>
			<td>MEDICON</td>
			<td>06.50.15</td>
			<td></td>
		</tr>
		<tr>
			<td>Video processor</td>
			<td>STORZ</td>
			<td>Image 1S connect II TC201</td>
			<td></td>
		</tr>
		<tr>
			<td>Yankauer suction tube</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

learning modern laryngeal surgery in a dissection laboratory

Surgery for laryngeal malignancies requires millimetric accuracy from the different endoscopic and open techniques available. Practice of this surgery is almost completely reserved to a few referral centers that deal with a large proportion of this pathology. Practice on human specimens is not always possible for ethical, economic, or availability reasons. The aim of this study is to provide a reproducible method for the organization of a laryngeal laboratory on ex vivo animal models where it is possible to approach, learn, and refine laryngeal techniques. Porcine and ovine larynges are ideal, affordable, models to simulate laryngeal surgery given their similarity to the human larynx in their anatomical layout and tissue composition. Herein, the surgical steps of transoral laser surgery, open partial horizontal laryngectomy, and total laryngectomy are reported. The merging of endoscopic and exoscopic views guarantees an inside-out perspective, which is vital for the comprehension of the complex laryngeal anatomy. The method was successfully adopted during three sessions of a dissection course &#34;Lary-Gym&#34;. Further perspectives on robotic surgical training are described.

This protocol proved to be useful for setting up a surgical training laboratory focused on laryngeal surgery using basic instruments and animal waste innards from the meat supply chain. The goal is mostly instructive, but it could be used by less experienced surgeons to improve their anatomical knowledge and surgical skills.
The protocol was adopted in three sessions of the authors&#39; dissection course organized in the &#39;Lary-Gym&#39; and in the second session of the Head and Neck Surgery Course named &#34;Better than live&#34;, where the laboratory dissections were accompanied by teaching sessions by skilled surgeons in this field, and it was greeted enthusiastically by the participants. Overall, 228 colleagues took part in both courses. Twenty-eight attended the Lary-Gym course, and 200 attended the &#39;Better than live&#39; course. In the Lary-Gym course&#39;s last two sessions, the satisfaction of 14 participants was determined through a dedicated questionnaire where participants responded to questions about their experience in the course. The questionnaire and the results are reported in Table 1. The animal models chosen proved to be very similar to the human counterpart, with a comparable tissue composition. The possibility to use both the endoscopic and open procedures guaranteed comprehensive understanding of the anatomical layout and surgical techniques. In fact, this inside-out vision could clarify the complex laryngeal anatomy and the implications of the surgical maneuvers in terms of extirpative and reconstructive procedures (e.g., the anastomosis technique in OPHL). In the last session of the course, human specimens and a surgical robot were successfully used to show various transoral robotic surgery (TORS) procedures. The setting of the room was similar to that described, showing that this protocol has good flexibility and can be adapted to equipment and space available in a particular institution.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60407/60407fig01.jpg" /> 
Figure 1: Endoscopic dissection. A young surgeon working in our endoscopic station on an animal specimen. <a href="https://www.jove.com/files/ftp_upload/60407/60407fig01large.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 colspan="2">Question</td>
			<td>1</td>
			<td>2</td>
			<td>3</td>
			<td>4</td>
			<td>5</td>
		</tr>
		<tr>
			<td colspan="2">How do you value the relevance of the treated topic in respect of your need to update your surgical skills?</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>1 (7%)</td>
			<td>13 (93%)</td>
		</tr>
		<tr>
			<td colspan="2">How do you value the educational quality of this course?</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>1 (7%)</td>
			<td>13 (93%)</td>
		</tr>
		<tr>
			<td colspan="2">How do you value the utility of this course?</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>1 (7%)</td>
			<td>13 (93%)</td>
		</tr>
		<tr>
			<td colspan="2">Absence of conflict of interest.</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>14 (100%)</td>
		</tr>
	</tbody>
</table>
Table 1: The Lary-Gym course: satisfaction questionnaire and responses. The score ranges from 1 (very dissatisfied) to 5 (very satisfied). Percentages are reported in brackets.

Watch this Scientific Journal Video about Learning Modern Laryngeal Surgery in a Dissection Laboratory at JoVE.com

Learning Modern Laryngeal Surgery in a Dissection Laboratory

The purpose of this paper is to illustrate how to organize a reproducible laboratory for laryngeal surgery on affordable and closely similar animal laryngeal models in order to improve anatomical and surgical knowledge and skills.

Surgery for laryngeal malignancies requires millimetric accuracy from the different endoscopic and open techniques available. Practice of this surgery is ...

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Medicine

7.8K Views.  Candiolo Cancer Institute, FPO-IRCCS. The purpose of this paper is to illustrate how to organize a reproducible laboratory for laryngeal surgery on affordable and closely similar animal laryngeal models in order to improve anatomical and surgical knowledge and skills.

Video: Learning Modern Laryngeal Surgery in a Dissection Laboratory

Discovering Middle Ear Anatomy by Transcanal Endoscopic Ear Surgery: A Dissection Manual

The middle ear is located in the center of the temporal bone and bears a highly complex anatomy. The recently introduced exclusively endoscopic transcanal approach to the middle ear is a minimally invasive technique sparing the bone and mucosa of the mastoid bone, since the middle ear is accessed through the external auditory canal. This emerging method has several advantages over the traditional (microscopic) approaches to the middle ear such as the panoramic wide-angle views of the anatomy, the possibility to approach and magnify tiny structures, and the possibility of looking around the corner using angled endoscopes.
The cadaveric dissection method presented here consists of an overview on the technical requirements and a precise description of a step-by-step protocol to discover the anatomy of the middle ear. Each step and anatomical structure is carefully described in order to provide a comprehensive guide to endoscopic ear anatomy. In our opinion, this is particularly important to any novice in endoscopic ear surgery as it provides thorough anatomical knowledge and may improve surgical skills.

The aim of this article is to describe the methodology of exclusively endoscopic cadaveric middle ear dissection. Moreover, we aim to provide a comprehensive guide to endoscopic middle ear anatomy.

The middle ear is located in the center of the temporal bone and bears a highly complex anatomy. The recently introduced exclusively endoscopic transcanal ...

Emergency Undocking in Robotic Surgery: A Simulation Curriculum

The following is a training platform to allow robotic surgeons to develop the skills necessary to lead an interprofessional team in emergency undocking of a robotic system. In traditional robotic training for surgeons, a brief web based overview of performing an emergency undocking is provided during initial introductory training to the robotics system. During such a process, there is no training in delineation of interdisciplinary roles for operating room (OR) personnel. The training presented here uses formative simulation and debriefing followed by a lecture. For the simulation, a modified gynecologic simulator is draped in a steep trendelenburg position consistent with most gynecologic laparoscopic surgery. The training torso is modified using tubing hooked to pressure bags of red food colored IV fluid used to simulate a catastrophic vessel injury on demand. Positioned throughout the operating room setting is an interprofessional team consisting of embedded standardized persons (ESPs) to fulfill the roles of a circulating nurse, scrub nurse, anesthesiologist, and bedside assist surgeon. Robotic surgeons are presented a case scenario necessitating emergency undocking, and given control of the robotic instruments. The scenario is terminated following either successful completion of an emergency undock, or at five minutes due to the emergent nature of the case. A debriefing session with hands on training of the steps to emergency undocking, necessary equipment, troubleshooting techniques, and operating room personnel roles follows the simulation. The learners are presented with a short lecture reemphasizing the material presented in the debriefing for their own self-study. This training results in improved time accessing the patient, improved knowledge, confidence, and completion of critical actions, and can be replicated in most institutions. All robotic surgeons should be able to demonstrate competence in this crucial intervention. A limitation of the curriculum is ability to access the in-situ environment for training purposes.

This training platform is designed to allow robotic surgeons to develop the skills necessary to lead an interprofessional team in emergency undocking of the robotic system. Training includes the utilization of the technology and equipment to perform an emergency undocking, as well as delineation of roles for such a scenario.

The following is a training platform to allow robotic surgeons to develop the skills necessary to lead an interprofessional team in emergency undocking of ...

Intra-Operative Neural Monitoring of Thyroid Surgery in a Porcine Model

Intraoperative injury to the recurrent laryngeal nerve (RLN) can cause vocal cord paralysis, which interferes with speech and can potentially interfere with breathing. In recent years, intraoperative neural monitoring (IONM) has been widely adapted as an adjunct technique to localize the RLN, detect RLN injury, and predict vocal cord function during the operations. Many studies have also used animal models to investigate new applications of IONM technology and to develop reliable strategies for preventing intraoperative RLN injury. The aim of this article is to introduce a standard protocol for using a porcine model in IONM research. The article demonstrates the procedures for inducing general anesthesia, performing tracheal intubation, and experimental design to investigate the electrophysiological characteristics of RLN injuries. Applications of this protocol can improve overall efficacy in implementing the 3R principle (replacement, reduction and refinement) in porcine IONM studies.

This study aims to develop a standard protocol of intra-operative neural monitoring of thyroid surgery in a porcine model. Here, we present a protocol to demonstrate general anesthesia, to compare different types of electrodes, and to investigate the electrophysiological characteristics of the normal and injured recurrent laryngeal nerves.

Intraoperative injury to the recurrent laryngeal nerve (RLN) can cause vocal cord paralysis, which interferes with speech and can potentially interfere ...

A Neonatal BALB/c Mouse Model of Necrotizing Enterocolitis

Necrotizing enterocolitis (NEC) is the most severe gastrointestinal (GI) disease that often occurs in premature infants, especially very low birth weight infants, with high mortality and unclear pathogenesis. The cause of NEC may be related to inflammatory immune regulatory system abnormalities. An NEC animal model is an indispensable tool for NEC disease immune research. NEC animal models usually use C57BL/6J neonatal mice; BALB/c neonatal mice are rarely used. Related studies have shown that when mice are infected, Th2 cell differentiation is predominant in BALB/c mice compared to C57BL/6J mice. Studies have suggested that the occurrence and development of NEC are associated with an increase in T helper type 2 (Th2) cells and are generally accompanied by infection. Therefore, this study used neonatal BALB/c mice to induce an NEC model with similar clinical characteristics and intestinal pathological changes as those observed in children with NEC. Further study is warranted to determine whether this animal model could be used to study Th2 cell responses in NEC.

Necrotizing enterocolitis (NEC) is the most severe gastrointestinal (GI) disease that often occurs in premature infants, especially very low birth weight infants, with high mortality and unclear pathogenesis. The cause of NEC may be related to inflammatory immune regulatory system abnormalities. An NEC animal model is an indispensable tool for NEC disease immune research. NEC animal models usually use C57BL/6J neonatal mice; BALB/c neonatal mice are rarely used. Related studies have shown that when mice are infected, Th2 cell differentiation is predominant in BALB/c mice compared to C57BL/6J mice. Studies have suggested that the occurrence and development of NEC are associated with an increase in T helper type 2 (Th2) cells and are generally accompanied by infection. Therefore, this study used neonatal BALB/c mice to induce an NEC model with similar clinical characteristics and intestinal pathological changes as those observed in children with NEC. Further study is warranted to determine whether this animal model could be used to study Th2 cell responses in NEC.

Necrotizing enterocolitis (NEC) is the most severe gastrointestinal (GI) disease that often occurs in premature infants, especially very low birth weight ...

Extrahepatic Bile Duct and Gall Bladder Dissection in Nine-Day-Old Mouse Neonates

The dissection of murine neonatal bile ducts has been described as difficult. The main aim of the described standard operating procedure is the isolation of the extrahepatic bile duct (EBD) in mouse neonates without damaging the bile duct during preparation. Because of its exceptionally close preparation compared to the cholangiocytes cell line and harvesting of the entire extrahepatic bile duct system (EBDS), the described approach is extremely useful in researching animal models of newborn bile duct disorders, such as biliary atresia. After euthanasia, the peritoneal cavity was accessed, and the bile duct system, duodenum, and liver were extracted with the unique En-bloc-Resection (EbR). The extracted sample is placed on a foam mat, and the EBD is dissected from contaminating cells atraumatically without necessary touch. The dissection of the entire EBDS is a significant advantage of this method. Caution must be taken due to the small size and amount of bile duct tissue. Using the described technique, there is no damage to the cholangiocytes. Further, the purity of the technique is reproducible (n = 10). Therefore, optimally comparable samples can be harvested. Furthermore, no bile duct tissue is harmed, because any contact with the bile duct system can be avoided during preparation, leaving the bile fluid inside the gall bladder. Most importantly, while performing the final gall bladder and bile duct dissection, atraumatic microinstruments were used only slightly lateral of the bile duct without squeezing it. This is the key to a clean and intact sample, and essential for further histological investigation or the isolation of cholangiocytes. To summarize, the described innovative dissection technique enables especially inexperienced operators with the necessary equipment to isolate the EBDS as cleanly as possible.

For the observation of murine neonatal bile duct disorders, an intact bile duct and efficient preparation are required. Therefore, a new approach for isolating the entire extrahepatic bile duct system in murine neonates was successfully developed while maintaining the integrity of the bile duct.

The dissection of murine neonatal bile ducts has been described as difficult. The main aim of the described standard operating procedure is the isolation ...

A Heterotopic Mouse Model for Studying Laryngeal Transplantation

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other animal models. Although first described by Shipchandler et al. in 2009, this technique is not widely used, possibly due to the difficulties in learning the microsurgical technique and time required to master it. This paper describes the surgical steps in detail, as well as potential pitfalls to avoid, in order to encourage effective use of this technique.
In this model, the bilateral carotid arteries of the donor larynx are anastomosed to the recipient carotid artery and external jugular vein, allowing for blood flow through the graft. Blood flow can be confirmed intraoperatively by the visualization of blood filling in the graft bilateral carotid arteries, reddening of the thyroid glands of the graft, and bleeding from micro vessels in the graft. The crucial elements for success include delicate preservation of the graft vessels, making the correct size arteriotomy and venotomy, and using the appropriate number of sutures on the arterial-arterial and arterial-venous anastomoses to secure vessels without leakage and prevent occlusion. 
Anyone can become proficient in this model with sufficient training and perform the procedure in approximately 3 h. If performed successfully, this model allows for immunologic studies to be performed with ease and at low cost.

The aim of this manuscript is to describe the microsurgical steps required to perform a heterotopic laryngeal transplant in mice. The advantages of this mouse model compared to other animal models of laryngeal transplantation are its cost-effectiveness and the availability of immunologic assays and data.

Laryngeal heterotopic transplantation, although a technically challenging procedure, offers more scientific analysis and cost benefits compared to other ...

Comparative Study of Blood Collection Techniques in Rats

Modified Tail Vein and Penile Vein Puncture for Blood Sampling in the Rat Model

Blood samples are required in most experimental animal designs to assess various hematological parameters. This paper presents two procedures for blood collection in rats: the lateral tail vein puncture and the dorsal penile vein puncture, which offer significant advantages over other previously described techniques. This study shows that these two procedures allow for fast sampling (under 10 min) and yield sufficient blood volumes for most assays (202 &#956;L &#177; 67.7 &#956;L). The dorsal penile vein puncture must be done under anesthesia, whereas the lateral tail vein puncture can be done on a conscious, restrained animal.
Alternating these two techniques, therefore, enables blood draw in any situation. While it is always recommended for an operator to be assisted during a procedure to ensure animal welfare, these techniques require only a single operator, unlike most blood sampling methods that require two. Moreover, whereas these previously described methods (e.g., jugular stick, subclavian vein blood draw) require extensive prior training to avoid harm to or death of the animal, tail vein and dorsal penile vein puncture are rarely fatal. For all these reasons, and according to the context (e.g., for studies including male rats, during the perioperative or immediate postoperative period, for animals with thin tail veins), both techniques can be used alternately to enable repeated blood draws.

Author Spotlight: Context Dependent Blood Collection Methods in Rats 

Here, we present a protocol to offer rapid, easy, and reliable blood collection alternatives for the rat model. We describe three different blood sampling methods according to the context: tail vein puncture under anesthesia or on a conscious animal, and dorsal penile vein puncture under anesthesia.

Blood samples are required in most experimental animal designs to assess various hematological parameters. This paper presents two procedures for blood ...

Laryngeal Mask Airway (LMA) Placement in a Neonatal Patient Simulator Using a Non-Inflatable Supraglottic Airway (SGA)

The effective delivery of positive pressure ventilation (PPV) can be challenging during neonatal resuscitation. Achieving a patent airway through an appropriate interface during neonatal resuscitation is critical for avoiding airway obstruction and leakage and optimizing access to PPV. Due to the complexity of face mask ventilation, providers have explored corrective steps. However, these methods are difficult to master and thus may present a risk for ventilation delay and/or interruptions at the critical time of resuscitation and the development of complications. In addition, neonatal endotracheal intubation is an invasive procedure that requires significant practice and training. The supraglottic airway (SGA) is a useful laryngeal mask airway (LMA) interface that decreases the time required to achieve a secure airway and reduces the need for endotracheal intubation. Despite the available evidence regarding its effectiveness, insufficient training and awareness limit SGA use in the real world, and frontline providers report low confidence in SGA placement. Here we provide a detailed description of SGA placement, the instruction of which requires only minimal training and leads to a short time to proficiency. Briefly, after the administration of initial ventilatory corrective steps in a neonatal manikin, a provider inserts a non-inflatable SGA into the larynx. This method allows for a single individual to provide effective delivery of PPV in a noninvasive manner without the need for expensive equipment such as video laryngoscopy. Instructors can easily teach this technique with ease and little cost in any clinical and research setting. This is also true for different income settings, including high-, middle-, and low-income countries.

Laryngeal Mask Airway LMA Placement in a Neonatal Patient Simulator Using a Non-Inflatable Supraglottic Airway SGA

This protocol is designed to describe a method to place and establish a supraglottic (alternative) airway during neonatal resuscitation. It can be used in situations where frontline providers cannot ventilate or endotracheally intubate neonates requiring resuscitation.

The effective delivery of positive pressure ventilation (PPV) can be challenging during neonatal resuscitation. Achieving a patent airway through an ...

Gasless Endoscopic Thyroidectomy via the Trans-Axillary Approach

For patients with early, low-risk papillary thyroid carcinoma, an increasing number are opting for endoscopic thyroid surgery due to its ability to achieve favorable therapeutic outcomes while maintaining excellent cosmetic results. Among the available endoscopic procedures, the Gasless Endoscopic Thyroidectomy Trans-axillary Approach (GETTA) has gained popularity among surgeons. This is attributed to its straightforward cavity construction, spacious operating area, precise visual field exposure, and manageable learning curve. However, few studies have provided detailed descriptions of the specific surgical steps involved in GETTA. Drawing from a synthesis of existing literature and our own clinical expertise, we present a comprehensive outline of the GETTA procedure. This process can be categorized into five distinct stages: positioning and incision planning; establishment of surgical cavities; identification and protection of the recurrent laryngeal nerve, inferior parathyroid gland, and central neck dissection; localization and preservation of the superior laryngeal nerve, superior parathyroid gland, and dissection of the thyroid's superior pole; transection of the thyroid isthmus followed by en bloc resection of the thyroid gland and central neck lymph nodes. The five-step approach of GETTA is easy to learn and can be adapted for resecting both benign and malignant thyroid and parathyroid diseases.

Gasless Endoscopic Thyroidectomy via the Trans-Axillary Approach

This protocol presents the procedure for conducting thyroidectomy using the gasless endoscopic thyroidectomy trans-axillary approach (GETTA).

For patients with early, low-risk papillary thyroid carcinoma, an increasing number are opting for endoscopic thyroid surgery due to its ability to ...

Validating Xanthatin-Keap1 Interaction with Docking and CETSA

Protein Target Prediction and Validation of Small Molecule Compound

Proteins are fundamental to human physiology, with their targets being crucial in research and drug development. The identification and validation of crucial protein targets have become integral to drug development. Molecular docking is a computational tool widely utilized to investigate protein-ligand binding, especially in the context of drug and protein target interactions. For the experimental verification of the binding and to access the binding of the drug and its target directly, the cellular thermal shift assay (CETSA) method is used. This study aimed to integrate molecular docking with CETSA to predict and validate interactions between drugs and vital protein targets. Specifically, we predicted the interaction between xanthatin and Keap1 protein as well as its binding mode through molecular docking analysis, followed by verification of the interaction using the CETSA assay. Our results demonstrated that xanthatin could establish hydrogen bonds with specific amino acid residues of Keap1 protein and reduce the thermostability of Keap1 protein, indicating that xanthatin could directly interact with Keap1 protein.

Author Spotlight: Streamlining Protein Target Prediction and Validation via Molecular Docking and CETSA

The experiment used here shows a method of molecular docking combined with cellular thermal shift assay to predict and validate the interaction between small molecules and protein targets.

Proteins are fundamental to human physiology, with their targets being crucial in research and drug development. The identification and validation of ...