Name: complete thymectomy in adult rats with non-invasive endotracheal intubation
Uploaded: 2014-12-29T13:00:00.000+00:00
Duration: 8 min 35 s
Description: Watch this Scientific Journal Video about Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation at JoVE.com

This work was supported by AI101263 from the National Institutes of Health (T.V.B), and by a grant from The Hartwell Foundation (M.L.M). M.L.M. is a member of the Duke Comprehensive Cancer Center.

NOTE: All experimental procedures involving the use of rats were done in accordance with protocols approved by the Animal Care and Use Committee of Duke University.
1. Preparation of the Tracheal Intubation Cannula
<ol>
	<li>Cut off the needle end of a 2 inch 14 G angiocathether needle with cutting pliers.</li>
	<li>Pinch the lumen of the needle closed with needle-nosed pliers.</li>
	<li>Cut the pinched section down to the edge of the open-lumen portion with cutting pliers, and then cut the edges on both sides of the remaining end at 30 - 45&#176; angle to the needle edge.</li>
	<li>Place the bottom 3 - 4 mm of the cannula into the open area of straight slip-joint pliers and pull upwards until a slight upward curve has been formed at the end. 
	NOTE: This will facilitate directing the catheter into the trachea (Figure 1).</li>
	<li>Use fine sandpaper to smooth down the edges at the end of the cannula.</li>
	<li>Return the blunted instrument into the angiocatheter tubing. Ensure that the tubing is slightly shorter than the blunted needle.</li>
</ol>
2. Pre-Surgical Procedures
<ol>
	<li>Set up the site for the procedure by placing an operating microscope set to 10X magnification over the operating area.</li>
	<li>Place a warming pad on the operating area and cover with a clean absorptive pad.</li>
	<li>Set up a heart rate and blood oxygenation monitor near the surgical field.</li>
	<li>Weigh the rat for weight-based medication dosing (e.g., postoperative analgesia, antibiotics, or lymphocyte depleting antibody). 
	NOTE: Typical adult rats weigh between 350 - 450 g.</li>
	<li>Sedate the rat with vaporized 3% isoflurane-O2 (3 L/min) using an induction chamber vented to a waste anesthetic gas scavenging system containing activated charcoal. Allow 5 min for anesthesia induction to provide deep anesthesia necessary for endotracheal intubation.</li>
	<li>Administer Carprofen or Meloxicane (4.4 mg/kg) subcutaneously to the rat prior to proceeding with intubation and surgery.</li>
	<li>Use electric clippers to shave the neck and chest of the anesthetized rat prior to intubation.</li>
</ol>
3. Intubation
<ol>
	<li>Prepare the intubation setup by rubbing a small amount of lubrication (e.g., K-Y gel) on the end of the intubation cannula. Turn on the ventilator to begin the flow of isoflurane gas through the connector tubing.</li>
	<li>Transfer the rat to the intubation apparatus and suspend the rat on the metal bar by its upper incisor teeth (Figure 2A). 
	NOTE: Certain intubation apparatuses may be purchased online. The model shown here is homemade using an acrylic sheet molded around sections of a 2 x 4 inch lumbar piece using a butane torch, and reinforcements are attached with chloroform. The bar (or metal wire) is attached by drilling holes over masking tape placed on the side of the acrylic to prevent splintering or cracking.</li>
	<li>Trans-illuminate the rat&#8217;s neck by positioning a flexible high-intensity light source 1 - 2 cm from the ventral surface of the neck (Figure 2B).</li>
	<li>Use a pair of student standard pattern forceps to pull the tongue gently upwards and to the side of the bottom teeth. Grip the tongue between the thumb and forefinger of one hand while placing the flat inside surface of one side of the forceps against the lower end of the tongue. Press ventrally to expose the epiglottis and aperture of the larynx (Figure 2C).</li>
	<li>Visualize the vocal cords and guide the upturned, blunted end of the intubation cannula anteriorly through the open glottis into the trachea until the hub of the angiocatheter touches the incisors.</li>
	<li>Remove the metal stylet, and attach the anesthesia tubing to the angiocatheter opening to begin ventilation with isoflurane. 
	NOTE: The typical oxygen flow rate to the ventilator is 3 L/min with 3% isoflurane.
	<ol>
		<li>Set the ventilator at a rate of 60 respirations/min in a volume controlled ventilator mode that achieves a pressure of ~12 - 14 mmHg. Use a positive end-expiratory pressure (PEEP) of 3 cm H2O.</li>
	</ol>
	</li>
	<li>Observe bilateral chest wall expansion in sync with the ventilator to ensure proper placement of the endotracheal tube.</li>
	<li>Apply veterinarian-recommended eye ointment to the rat&#8217;s eyes to prevent dryness while under anesthesia.</li>
	<li>Apply veterinarian ointment on the rat&#8217;s eyes to prevent dryness while under anesthesia.</li>
	<li>Secure the endotracheal tube to the head of the rat with a strip of cloth tape. Ensure a firm connection between the endotracheal tube and the anesthetic inflow tubing.</li>
	<li>Attach the blood oxygenation and heart rate monitor to the rat&#8217;s foot and begin monitoring. Confirm proper anesthetization by confirming there is no reaction to toe-pinch.</li>
</ol>
4. Thoracotomy and Thymectomy
<ol>
	<li>Follow standard aseptic technique for the entirety of the procedure.
	<ol>
		<li>Clean the working area and operating table and disinfect with a 70% ethanol solution.</li>
		<li>Use sterile surgical gloves during the procedure, and autoclave all instruments and materials used during the procedure.</li>
		<li>Apply providone-iodine to the entire chest area and allow it to dry. Then clean the surface of the skin with 70% ethanol on gauze. Cover the rat with clear plastic wrap cutting a hole to expose the sterile operating field.</li>
	</ol>
	</li>
	<li>Identify the suprasternal notch in the upper thoracic region. Make a 2 cm midline incision through the skin starting 2 - 3 mm above the suprasternal notch and extending distally midline along the sternum using blunt-tipped Shea scissors.</li>
	<li>Perform a 1.5 cm&#160;median sternotomy from the suprasternal notch again using blunt-tipped Shea scissors. Keep the bottom edge of the scissors just underneath the sternum and advance slowly.</li>
	<li>Insert a small Alm retractor just underneath the separated sternum and open to reveal the pre-tracheal strap muscles (sternohyoid and sternothyroid muscles). Separate the pre-tracheal strap muscles using blunt Graefe forceps.&#160; NOTE: At this point, the trachea can be seen, and the intubation tube should be visualized inside the trachea.</li>
	<li>Place the prongs of the&#160;small Alm retractor underneath the separated strap muscles and sternum. Open the retractor to expose the superior aspect of the thymus.</li>
	<li>Use fine Dumont forceps to release the lateral edges of the thymus tissue and expose the lower thymic lobes.</li>
	<li>Pull the thymus gently superiorly into the open incision site, taking care to avoid contact with the superior vena cava, subclavian and carotid vessels, and to minimize disruption of the delicate pleural lining between the thymus and the lungs.</li>
	<li>As the thymic vessels are dissected and revealed, use micro-scissors to divide them. Use a cotton swab to hold pressure on the vessels for hemostasis if needed.</li>
	<li>Deliver the lower thymic lobes into the incision and then sharply lyse the posterior attachments. Remove the intact thymus and carefully inspect the removed thymus for any missing sections. 
	NOTE: Small thoracic lymph nodes may be visualized surrounding the thymus and are often difficult to differentiate from thymic tissue. These will be discrete round nodes with a similar appearance to thymic tissue but will not be in continuity with the thymus.</li>
	<li>Remove the Alm retractor and close the sternohyoid and sternothyoid muscles with two interrupted 5-0 Maxon sutures.</li>
	<li>Apply 2 drops of surgical grade cyanoacrylate tissue adhesive across the sutures to seal the mediastinum under positive pressure ventilation. 
	NOTE: This will decrease the incidence of pneumothorax and hematoma.</li>
	<li>Close the sternum with two interrupted 4-0 silk sutures on a cutting needle. Insert the needle through gaps between the ribs, taking care to guide the needle just under the sternum to avoid the underlying muscle layer.</li>
	<li>Close the skin layer with a running 4-0 Nylon suture. Discontinue the isoflurane at this point to shorten the post-operative anesthesia recovery period.</li>
	<li>Clean the incision site and surrounding skin with saline-moistened gauze.</li>
	<li>Apply several drops of Bupivicaine (0.25%) for local anesthesia, followed by 1 - 2 drops of cyanoacrylate glue over the incision to seal the incision.</li>
	<li>Continue to ventilate until the rat shows signs of independent respiratory effort and begins to move its extremities. Extubate the rat and allow it to recover under close observation in a cage placed on a warming pad.</li>
	<li>Administer Buprenorphine (0.01 - 0.05 mg/kg) subcutaneously to the rat immediately post-operatively and repeat every 8-12 hr&#160;for 48 hr&#160;post-operatively (normally 5 total doses). At 24 and 48 hr&#160;post-operatively, administer Carprofen or Meloxicane (4.4 mg/kg) subcutaneously (3 total doses including the pre-operative dose).</li>
	<li>Continue to monitor until the rat is moving around the cage. At any signs of respiratory distress, perform re-intubation and re-exploration of the chest.
	<ol>
		<li>Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbency.</li>
		<li>Do not return an animal that has undergone surgery to the company of other animals until fully recovered.</li>
	</ol>
	</li>
	<li>Remove the skin layer sutures 1 week post-operatively.</li>
</ol>

<ol>
	<li>Hard, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thymectomy+in+the+neonatal+rat.">Thymectomy in the neonatal rat.</a> Lab. Anim. 9 (2), 105-110 (1975).</li><li>Kobayashi, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+technique+for+complete+thymectomy+in+adult+rats.">A technique for complete thymectomy in adult rats.</a> J. Immunol. Methods. 1759 (94), 0 (1994).</li><li>Delrivi&#232;re, L., Gibbs, P., Kobayashi, E., Kamada, N., Gianello, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+technique+of+complete+thymectomy+in+adult+rats+without+tracheal+intubation.">New technique of complete thymectomy in adult rats without tracheal intubation.</a> Microsurgery. 18 (1), 6-8 (1998).</li><li>Walker, K. G., Jacques, B. C., Tweedle, J. R., Bradley, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simpler+technique+for+open+thymectomy+in+adult+rats.">A simpler technique for open thymectomy in adult rats.</a> J. Immunol. Methods. 175 (1), 141 (1994).</li><li>Na, N., Zhao, D., Huang, Z., Hong, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+improved+method+for+rat+intubation+and+thymectomy.">An improved method for rat intubation and thymectomy.</a> Chin. Med. J. (Engl). 124 (17), 2723-2727 (2011).</li><li>Kastl, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simplification+of+rat+intubation+on+inclined+metal+plate).">Simplification of rat intubation on inclined metal plate).</a> Adv. Physiol. Educ. 28 (1-4), 29-34 (2004).</li><li>Weksler, B., Ng, B., Lenert, J., Burt, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simplified+method+for+endotracheal+intubation+in+the+rat.">A simplified method for endotracheal intubation in the rat.</a> J. Appl. Physiol. 76 (4), 1823-1825 (1994).</li><li>Jou, I. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simplified+rat+intubation+using+a+new+oropharyngeal+intubation+wedge.">Simplified rat intubation using a new oropharyngeal intubation wedge.</a> J. Appl. Physiol. 89 (5), 1766-1770 (2000).</li><li>Rivard, A. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+intubation+and+ventilation+for+surgical+research.">Rat intubation and ventilation for surgical research.</a> J. Invest. Surg. 19 (4), 267-274 (2006).</li><li>Zhao, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+model+of+isolated,+vascular+whole+thymus+transplantation+in+nude+rats.">A model of isolated, vascular whole thymus transplantation in nude rats.</a> Transplant Proc. 44 (5), 1394-1398 (2012).</li><li>AbuAttieh, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Affinity+maturation+of+antibodies+requires+integrity+of+the+adult+thymus.">Affinity maturation of antibodies requires integrity of the adult thymus.</a> Eur. J. Immunol. 42 (2), 500-510 (2012).</li><li>Reeves, J. P., Reeves, P. A., Chin, L. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival+surgery:+removal+of+the+spleen+or+thymus.">Survival surgery: removal of the spleen or thymus.</a> Curr. Protoc. Immunol. 1 (Unit 1.10), (2001).</li><li>Siemionow, M., Izycki, D., Ozer, K., Ozmen, S., Klimczak, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+thymus+in+operational+tolerance+induction+in+limb+allograft+transplant+model.">Role of thymus in operational tolerance induction in limb allograft transplant model.</a> Transplantation. 81 (11), 1568-1576 (2006).</li><li>Satoh, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunosuppressive+effect+of+long-term+drainage+of+thoracic+duct+on+immunological+memory+in+adult+thymectomized+rats.+Transplant.">Immunosuppressive effect of long-term drainage of thoracic duct on immunological memory in adult thymectomized rats. Transplant.</a> Proc. 37 (4), 1947-1948 (2005).</li><li>Groen, H., Klatter, F., Pater, J., Nieuwenhuis, P., Rozing, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporary,+but+Essential+Requirement+of+CD8++T+Cells+Early+in+the+Pathogenesis+of+Diabetes+in+BB+Rats+as+Revealed+by+Thymectomy+and+CD8+Depletion.">Temporary, but Essential Requirement of CD8+ T Cells Early in the Pathogenesis of Diabetes in BB Rats as Revealed by Thymectomy and CD8 Depletion.</a> Clin. Dev. Immunol. 10 (2-4), 141-151 (2003).</li></ol>

The authors have nothing to disclose.

The present protocol for complete thymectomy provides a minimally invasive approach with a three layer incision closure with surgical glue application designed to minimize complications. Complete removal of the thymus was demonstrated by the loss of na&#239;ve T cells and by IHC staining of mediastinal lymphoid tissue for cytokeratin.
The procedure of adult rat thymectomy has been complicated by mortality rates ranging from 1.5 - 20% due to peri-operative complications, most of which are the a...

Since the early 1960s, the thymus has been recognized for its critical role in the development of central immunological tolerance. Rodent thymectomy has proven to be an essential procedure in defining the role of the thymus in lymphocyte differentiation, self-tolerance, and immunotolerance in the setting of allograft transplantation and tumor metastasis. Removal of the rat thymus allows for studies involving T-cell depletion or the adoptive transfer of defined T cell populations without the re-emergence of native na&#239;ve T cells.
Thymectomies in neonatal rodents can be accomplished using a suction technique with reliable outcomes1. In adult rats, this technique is associated with an approximate 20% mortality rate and frequently results in an incomplete thymectomy2. To consistently achieve complete thymectomy in adult rats, open exposure of the mediastinum through a median sternotomy is required. However, this procedure is associated with complications that include tracheal injury, hemorrhage and pneumothorax leading to an overall mortality rate ranging from 1.5 - 6%2-4.
Over the last two decades improvements in thymectomy techniques have decreased perioperative complications and have improved survival rates. Endotracheal intubation allowing for positive pressure ventilation has lessened pneumothorax rates5. Methods of intubation previously described range from open exposure to the trachea to less invasive methods using direct vocal cord visualization. Complications associated with the intubation procedure include tracheal injury, vocal cord rupture, unintended esophageal intubation, and hemorrhage resulting from cardiac puncture or laceration of the superior vena cava. In addition, close proximity of the lower thymic lobes to the pleural lining can result in pneumothorax.
Here we describe a technique of thymectomy through a minimally invasive 2 cm skin incision following a simple method of endotracheal intubation using a blunted-end angiocatheter and transcutaneous tracheal illumination. The thymectomy involves a 1.5&#160;cm sternotomy and a three-layer closure with surgical glue application to seal the mediastinum and minimize the incidence of hemorrhage and respiratory complications. This method reliably results in complete thymectomy as evidenced by the disappearance of CD4+ and CD8+ na&#239;ve T cells following thymectomy and the absence of thymic tissue on IHC staining. Operative times and peri-procedural mortality are kept to a minimum.

<table><tbody><tr><td>2 inch 14 G angiocatheter</td><td></td><td></td><td></td></tr><tr><td>Operating microscope</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Warming pad</td><td></td><td></td><td></td></tr><tr><td>Heart rate and blood oxygenation monitor for rodents with foot sensors</td><td>Harvard Apparatus</td><td>ST1 72-8010, ST1 72-8098 (Rat foot sensor)</td><td></td></tr><tr><td>Intubation apparatus (plastic with metal bar at the top)</td><td></td><td></td><td>See &lt;strong&gt;Figure 2&lt;/strong&gt;</td></tr><tr><td>Small animal anesthesia system with induction box, isoflurane tank and O&lt;sub&gt;2&lt;/sub&gt; tank</td><td>Harvard Apparatus</td><td>ST1 72-6420</td><td></td></tr><tr><td>Small animal ventilator with tubing</td><td>CWE</td><td>12-02000 (ventilator) and 12-04000 (external valve assembly for mice/rats)</td><td></td></tr><tr><td>High-intensity fiber optic Illuminator</td><td>Dolan Jenner</td><td>EEG 2823M</td><td></td></tr><tr><td>Student standard pattern forceps</td><td>Fine Science Tools</td><td>91100-16</td><td></td></tr><tr><td>Fine straight scissors</td><td>Fine Science Tools</td><td>14060-09</td><td></td></tr><tr><td>Blunt-tipped Shea scissors</td><td>Fine Science Tools</td><td>14105-12</td><td></td></tr><tr><td>Small Alm retractor (for sternum)</td><td>Fine Science Tools</td><td>17008-07</td><td></td></tr><tr><td>Blunt Graefe forceps</td><td>Fine Science Tools</td><td>11050-10</td><td></td></tr><tr><td>Fine Dumont forceps</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>5-0 Maxon sutures</td><td>Ethicon</td><td></td><td></td></tr><tr><td>4-0 Silk sutures (with cutting needle)</td><td>Ethicon</td><td></td><td></td></tr><tr><td>6-0 Nylon suture</td><td>Ethicon</td><td></td><td></td></tr><tr><td>Cyanoacrylate glue (Endermil)</td><td></td><td></td><td></td></tr><tr><td>Lubrication gel</td><td>Akorn Animal Health</td><td>NDC 17478-162-35</td><td></td></tr></tbody></table>

complete thymectomy in adult rats with non-invasive endotracheal intubation

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and pneumothorax from pleural disruption can result in a significant mortality rate. This protocol is a simple method of rat thymectomy that utilizes a mini-sternotomy and endotracheal intubation. Intubation is accomplished with a non-invasive and easily reproducible method and allows for positive pressure ventilation to prevent pneumothorax and a controlled airway that allows sufficient time for careful thymus dissection to minimize pleural disruption. A 1.5 cm sternal incision decreases contact with mediastinal vessels and pleura, while still providing full visualization of the thymus. Following exposure of the mediastinum, the thymus is removed by blunt dissection under magnification. The pleural space is then sealed by suture closure of the pre-tracheal muscles followed by the application of surgical glue. The thorax is then closed by suture closure of the sternum, followed by suture closure of the skin. All thymectomies were complete as evidenced by immunohistochemical (IHC) staining of mediastinal tissue, and absence of na&#239;ve T-cells by flow cytometry, and the procedure had a 96% survival rate. This method is suitable when complete thymectomy with minimal complications is desired for further immunological studies in athymic adult rats.

This procedure was performed on adult Lewis rats (n = 26). The mean operation time was 15 &#177; 3 min. There was no intraoperative mortality. The mean intubation time &#8211; from placement of rat on the intubation apparatus to establishment of ventilation &#8211; was 45 &#177; 5 sec. 24 rats had a full recovery from the operation with no evidence of respiratory difficulties or hemorrhage through post-operative day (POD) 14. One rat developed respiratory difficulty on POD 4 and underwent re-operation to explore the medi...

Watch this Scientific Journal Video about Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation at JoVE.com

Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described.

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and ...

complete-thymectomy-adult-rats-with-non-invasive-endotracheal

Research

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Immunology and Infection

16.5K Views.  Duke University Medical Center. Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described. 

Video: Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are necessary for whole animal experiments that require surgery under deep anesthesia or measurements of lung function. Tracheostomy has been the standard for intubating the airway in these mice to allow mechanical ventilation. Orotracheal intubation has been reported but has not been successfully used in many studies because of the substantial technical difficulty or a requirement for highly specialized and expensive equipment. Here we report a technique of direct laryngoscopy using an otoscope fitted with a 2.0 mm speculum and using a 20 G&#160;intravenous catheter as an endotracheal tube. We have used this technique extensively and reliably to intubate and conduct accurate assessments of lung function in mice. This technique has proven safe, with essentially no animal loss in experienced hands. Moreover, this technique can be used for repeated studies of mice in chronic models.

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

We have developed a simple, reliable, and relatively inexpensive method for endotracheal intubation in mice via direct laryngoscopy using an otoscope with a 2.0 mm speculum. This technique is atraumatic and can be used for repeated measurements in chronic experiments. We find it superior to tracheostomy or previously reported nonsurgical techniques.

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are ...

Medicine

Automated Measurement of Microcirculatory Blood Flow Velocity in Pulmonary Metastases of Rats

Because the lung is a major target organ of metastatic disease, animal models to study the physiology of pulmonary metastases are of great importance. However, very few methods exist to date to investigate lung metastases in a dynamic fashion at the microcirculatory level, due to the difficulty to access the lung with a microscope. Here, an intravital microscopy method is presented to functionally image and quantify the microcirculation of superficial pulmonary metastases in rats, using a closed-chest pulmonary window and automated analysis of blood flow velocity and direction. The utility of this method is demonstrated to measure increases in blood flow velocity in response to pharmacological intervention, and to image the well-known tortuous vasculature of solid tumors. This is the first demonstration of intravital microscopy on pulmonary metastases in a closed-chest model. Because of its minimized invasiveness, as well as due to its relative ease and practicality, this technology has the potential to experience widespread use in laboratories that specialize on pulmonary tumor research.

A method is presented to measure microcirculatory blood flow velocity in pulmonary cancer metastases of the pleural surface in rats in an automated fashion, using closed-chest pulmonary intravital microscopy. This model has potential to be used as a widespread tool to perform physiologic research on pulmonary metastases in rodents.

Because the lung is a major target organ of metastatic disease, animal models to study the physiology of pulmonary metastases are of great importance. ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Invasive Hemodynamic Characterization of the Portal-hypertensive Syndrome in Cirrhotic Rats

This is a detailed protocol describing invasive hemodynamic measurements in cirrhotic rats for the characterization of portal hypertensive syndrome. Portal hypertension (PHT) due to cirrhosis is responsible for the most severe complications in patients with liver disease. The full picture of the portal hypertensive syndrome is characterized by increased portal pressure (PP) due to the increased intrahepatic vascular resistance (IHVR), hyperdynamic circulation, and increased splanchnic blood flow. Progressive splanchnic arterial vasodilation and increased cardiac output with elevated heart rate (HR) but low arterial pressure characterizes the portal hypertensive syndrome.
Novel therapies are currently being developed that aim to decrease PP by either targeting IHVR or increased splanchnic blood flow &#8212; but side effects on systemic hemodynamics may occur. Thus, a detailed characterization of portal venous, splanchnic, and systemic hemodynamic parameters, including measurement of PP, portal venous blood flow (PVBF), mesenteric arterial blood flow, mean arterial pressure (MAP), and HR is needed for preclinical evaluation of the efficacy of novel treatments for PHT. Our video article provides the reader with a structured protocol for performing invasive hemodynamic measurements in cirrhotic rats. In particular, we describe the catheterization of the femoral artery and the portal vein via an ileocolic vein and the measurement of portal venous and splanchnic blood flow via perivascular Doppler-ultrasound flow probes. Representative results of different rat models of PHT are shown.

Here we describe a detailed protocol for invasive measurements of hemodynamic parameters including portal pressure, splanchnic blood flow, and systemic hemodynamics in order to characterize the portal hypertensive syndrome in rats.

This is a detailed protocol describing invasive hemodynamic measurements in cirrhotic rats for the characterization of portal hypertensive syndrome. ...

The Left Pneumonectomy Combined with Monocrotaline or Sugen as a Model of Pulmonary Hypertension in Rats

In this protocol, we detail the correct procedural steps and necessary precautions to successfully perform a left pneumonectomy and induce PAH in rats with the additional administration of monocrotaline (MCT) or SU5416 (Sugen). We also compare these two models to other PAH models commonly used in research. In the last few years, the focus of animal PAH models has moved towards studying the mechanism of angioproliferation of plexiform lesions, in which the role of increased pulmonary blood flow is considered as an important trigger in the development of severe pulmonary vascular remodeling. One of the most promising rodent models of increased pulmonary flow is the unilateral left pneumonectomy combined with a "second hit" of MCT or Sugen. The removal of the left lung leads to increased and turbulent pulmonary blood flow and vascular remodeling. Currently, there is no detailed procedure of the pneumonectomy surgery in rats. This article details a step-by-step protocol of the pneumonectomy surgical procedure and post-operative care in male Sprague-Dawley rats. Briefly, the animal is anesthetized and the chest is opened. Once the left pulmonary artery, pulmonary vein, and bronchus are visualized, they are ligated and the left lung is removed. The chest then closed and the animal recovered. Blood is forced to circulate only on the right lung. This increased vascular pressure leads to a progressive remodeling and occlusion of small pulmonary arteries. The second hit of MCT or Sugen is used one week post-surgery to induce endothelial dysfunction. The combination of increased blood flow in the lung and endothelial dysfunction produces severe PAH. The primary limitation of this procedure is that it requires general surgical skills.

The rodent left pneumonectomy is a valuable technique in pulmonary hypertension research. Here, we present a protocol to describe the rat pneumonectomy procedure and postoperative care to ensure minimal morbidity and mortality.

In this protocol, we detail the correct procedural steps and necessary precautions to successfully perform a left pneumonectomy and induce PAH in rats ...

A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR permits the experimental study of this specific process with continued alveolar ventilation, thereby avoiding the compounding injurious processes of hypoxia and atelectasis. In the clinical context, lung ischemia reperfusion injury (also known as lung IRI or LIRI) is caused by numerous processes, including but not limited to pulmonary embolism, resuscitated hemorrhagic trauma, and lung transplantation. There are currently limited effective treatment options for LIRI. Here, we present a reversible surgical model of lung IR involving first orotracheal intubation followed by unilateral left lung ischemia and reperfusion with preserved alveolar ventilation or gas exchange. Mice undergo a left thoracotomy, through which the left pulmonary artery is exposed, visualized, isolated, and compressed using a reversible slipknot. The surgical incision is then closed during the ischemic period, and the animal is awakened and extubated. With the mouse spontaneously breathing, reperfusion is established by releasing the slipknot around the pulmonary artery. This clinically relevant survival model permits the evaluation of lung IR injury, the resolution phase, downstream effects on lung function, as well as two-hit models involving experimental pneumonia. While technically challenging, this model can be mastered over the course of a few weeks to months with an eventual survival or success rate of 80%-90%.

A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR ...

Repeated Orotracheal Intubation in Mice

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ventral neck for direct confirmation of cannula placement in the trachea. The relative difficulty or the tissue trauma induced to the subject by such procedures can be an impediment to an investigator&#8217;s ability to perform longitudinal studies. This article illustrates a technique in which physical manipulation of the mouse following the use of a depilatory to remove hair from the ventral neck permits transcutaneous visualization of the trachea for orotracheal intubation regardless of degree of skin pigmentation. This method is innocuous to the subject and easily achieved with a limited understanding of murine anatomy. This refined approach facilitates repeated intubation, which may be necessary for monitoring progression of disease or instillation of treatments. Using this method may result in a reduction of the number of animals and technical skill required to measure lung function in mouse models of respiratory disease.

The goal of this article is to describe a refined method of intubation of the laboratory mouse. The method is noninvasive and, therefore, ideal for studies that require serial monitoring of respiratory function and/or instillation of treatments into the lung.

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ...

Exploring Obliterative Airway Disease in Lung Transplants

Murine Intrapulmonary Tracheal Transplantation: A Model for Investigating Obliterative Airway Disease After Lung Transplantation

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially reported by our team, this model has gained use in the study of OAD due to its high technical reproducibility and suitability for investigating immunological behaviors and therapeutic interventions.
In the IPTT model, a rodent tracheal graft is directly inserted into the recipient's lung through the pleura. This model is distinct from the heterotopic tracheal transplantation (HTT) model, wherein grafts are transplanted into subcutaneous or omental sites, and from the orthotopic tracheal transplantation (OTT) model in which the donor trachea replaces the recipient's trachea. 
Successful implementation of the IPTT model requires advanced anesthetic and surgical skills. Anesthetic skills include endotracheal intubation of the recipient, setting appropriate ventilatory parameters, and appropriately timed extubation after recovery from anesthesia. Surgical skills are essential for precise graft placement within the lung and for ensuring effective sealing of the visceral pleura to prevent air leakage and bleeding. In general, the learning process takes approximately 2 months.
In contrast to the HTT and OTT models, in the IPTT model, the allograft airway develops airway obliteration in the relevant lung microenvironment. This allows investigators to study lung-specific immunological and angiogenic processes involved in airway obliteration after lung transplantation. Furthermore, this model is also unique in that it exhibits tertiary lymphoid organs (TLOs), which are also seen in human lung allografts. TLOs are comprised of T and B cell populations and characterized by the presence of high endothelial venules that direct immune cell recruitment; therefore, they are likely to play a crucial role in graft acceptance and rejection. We conclude that the IPTT model is a useful tool for studying intrapulmonary immune and profibrotic pathways involved in the development of airway obliteration in the lung transplant allograft.

Author Spotlight: Investigating the Key Factors of Obliterative Bronchiolitis After Lung Transplantation

The murine intrapulmonary tracheal transplantation (IPTT) model is valuable for studying obliterative airway disease (OAD) after lung transplantation. It offers insights into lung-specific immunological and angiogenic behavior in airway obliteration after allotransplantation with high reproducibility. Here, we describe the IPTT procedure and its expected results.

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially ...

Tracheotomy-Based Endotracheal Intubation Protocol for Non-Survival Rat Models

Endotracheal Intubation via Tracheotomy and Subsequent Thoracotomy in Rats for Non-Survival Applications

Endotracheal intubation and subsequent ventilation are often basic requirements for translational research in rat models for various interventions that require controlled or high ventilation pressures or access to the thoracic cavity and organs. Conventional endoorotracheal intubation using the anatomically existing route through the mouth is well suited for survival experiments. However, this procedure poses some challenges, including generally higher levels of the required experience and technical skill, more advanced equipment, and greater time effort with relevant intubation failure rates and complications such as tracheal perforation, temporary systemic hypooxygenation, and relevant aerial leakage.
This manuscript, therefore, presents a detailed step-by-step protocol for endotracheal intubation through tracheotomy in non-survival rat models when guaranteed intubation success, constant oxygenation levels, high ventilation pressures, or open thoracotomy are required.
The protocol emphasizes the importance of meticulous surgical technique to ensure consistent and reliable outcomes, especially for researchers who are inexperienced or lack routine in the technique of endoorotracheal intubation via direct laryngoscopy. This procedure is, therefore, expected to minimize animal suffering and unnecessary animal losses.

Here we introduce a standardized procedure for endotracheal intubation via tracheotomy followed by thoracotomy in rats, aimed at enhancing the precision and reproducibility of non-survival applications requiring invasive ventilation and exposure of thoracic organs in in vivo rat models.

Endotracheal intubation and subsequent ventilation are often basic requirements for translational research in rat models for various interventions that ...

Optimizing Pulmonary Artery Banding for Consistent Right Heart Failure Induction

Rat Model of Right-Sided Cardiac Remodeling and Arrhythmia Using Pulmonary Artery Banding

Clinical conditions, including chronic obstructive pulmonary disease or pulmonary arterial hypertension (PAH), can lead to chronic right ventricle pressure overload and progressive right heart failure (RHF). RHF can be identified by right-sided cardiac hypertrophy and dilation associated with abnormal myocardial function affecting the RV and the right atrium (RA). We recently demonstrated that severe RHF is accompanied by an increased risk of atrial inflammation, atrial fibrosis, and atrial fibrillation (AF), the most common type of cardiac arrhythmia (CA). Recent studies have shown that RV and RA inflammation plays an important role in the arrhythmogenesis of CA, including AF. However, the impact of inflammation in the development of CA and AF in RHF is poorly described.
Experimental models of RHF are required to better understand the association between right-sided myocardial inflammation and CA. The rat model of monocrotaline (MCT)-induced pulmonary hypertension (PH) is well-established to provoke RHF. However, MCT triggers severe pneumo-toxicity and pulmonary inflammation. Hence, MCT-induced RHF does not help to distinguish whether the subsequent myocardial inflammation originates from the RHF per se or circulating inflammatory signals secreted by the injured lung.
In this article, a mechanical method involving pulmonary artery trunk banding (PAB) was used to provoke right-sided cardiac arrhythmogenesis. The PAB consists of performing a permanent suture of the pulmonary artery trunk for 3 weeks. Such an approach generates increased right-sided pressure overload. At D21 post-PAB, the suture results in hypertrophied, dilated, and inflamed RV and RA. The PAB-induced RHF is also accompanied by vulnerability to ventricular and atrial arrhythmias, including AF.

Right heart failure (RHF) is characterized by right-sided cardiac dilation and hypertrophy, leading to ventricular and atrial malfunction. Cardiopulmonary conditions associated with RHF are accompanied by an increased risk for cardiac arrhythmias. This article describes a standardized model of pulmonary artery banding-induced RHF associated with enhanced ventricular and atrial arrhythmogenesis.

Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

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Chapters in this video

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Automated Measurement of Microcirculatory Blood Flow Velocity in Pulmonary Metastases of Rats

An Improved Method for Rapid Intubation of the Trachea in Mice

Invasive Hemodynamic Characterization of the Portal-hypertensive Syndrome in Cirrhotic Rats

The Left Pneumonectomy Combined with Monocrotaline or Sugen as a Model of Pulmonary Hypertension in Rats

A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Repeated Orotracheal Intubation in Mice

Murine Intrapulmonary Tracheal Transplantation: A Model for Investigating Obliterative Airway Disease After Lung Transplantation

Endotracheal Intubation via Tracheotomy and Subsequent Thoracotomy in Rats for Non-Survival Applications

Rat Model of Right-Sided Cardiac Remodeling and Arrhythmia Using Pulmonary Artery Banding