The authors gratefully acknowledge the data storage service SDS@hd supported by the Ministry of Science, Research and the Arts Baden-W&#252;rttemberg (MWK) and the German Research Foundation (DFG) through grant INST 35/1314-1 FUGG and INST 35/1503-1 FUGG. Furthermore, the authors gratefully acknowledge the support from the NCT (National Center for Tumor Diseases in Heidelberg, Germany) through its structured postdoc program and the Surgical Oncology program. We also acknowledge the support through state funds approved by the State Parliament of Baden-W&#252;rttemberg for the Innovation Campus Health + Life Science Alliance Heidelberg Mannheim from the structured postdoc program for Alexander Studier-Fischer: Artificial Intelligence in Health (AIH) - A collaboration of DKFZ, EMBL, Heidelberg University, Heidelberg University Hospital, University Hospital Mannheim, Central Institute of Mental Health, and the Max Planck Institute for Medical Research. Furthermore, we acknowledge the support through the DKFZ Hector Cancer Institute at the University Medical Center Mannheim. For the publication fee, we acknowledge financial support by Deutsche Forschungsgemeinschaft within the funding program &#34;Open Access Publikationskoste&#34; as well as by Heidelberg University.

All animal activities described here have been approved by the institutional animal care and use committee (IACUC) of the Baden-W&#252;rttemberg Regional Council in Karlsruhe, Germany. Experimental animals were managed as per institutional standards and according to German laws for animal use and care and according to the directives of the European Community Council (2010/63/EU) as well as the ARRIVE guidelines and were conducted in accredited facilities. Male Spraque Dawley rats from Janvier Labs with an order weight of 400 g were used.
1. Procedure preparation (Figure 1)
<ol>
	<li>Prepare surgical preparation hooks by folding cannulas at an angle of 135&#176; at 1 cm to the tip, connecting them to plastic perfusion tubes via Luer-lock, and applying tension using a surgical mosquito clamp (Figure 1A,B).</li>
	<li>Prepare a rodent surgical exposure apparatus. 
	NOTE: Besides the availability of commercial platforms, these can also easily be self-constructed cost-effectively, as in this study.
	<ol>
		<li>For this, use a 40 cm x 50 cm steel plate with a 4 mm thickness. Create several 8 mm holes using a steel drill at the desired location for subsequent tension points. Place the Y-shaped fixation rods in these locations.</li>
	</ol>
	</li>
	<li>Place a heating pad on top of the steel plate of the surgical exposure apparatus in order to provide thermal support.</li>
	<li>Prepare the surgical preparation hooks with attached plastic tubes and surgical mosquito clamps to later apply tension to the tissue for surgical exposure.</li>
	<li>Remove the plug of a 10 mL plastic syringe with a male Luer-lock end and cut off the proximal end of the syringe. Insert the distal part into the ventilation bag as depicted and seal it airtight by covering it with a heat-shrinking tube and applying gentle heat using a laboratory torch (Figure 1D-F).</li>
	<li>Shorten and bevel different sizes of intravenous catheters and insert a guide wire from an arterial leader-cath set with the flexible wire end at the tip into the catheter size most appropriate for the respective animal size (presumably 14 G) for subsequent Seldinger technique22,23 during intubation (Figure 1G,H).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66684/66684fig01v2.jpg" /> 
Figure 1: Experimental setup. (A) A folded cannula connected to a perfusion tube used as a surgical preparation hook. (B) Preparation hook with clamp positioned in a metal fixation rod. (C) Sedated animal prior to intervention. (D) Ventilation bag from neonatology. (E-F) Airtight mounting of a Luer-lock syringe onto the ventilation bag using a heat-shrinking tube. (G) Shortened and beveled intravenous catheters in different sizes with a guide wire from an arterial leader-cath set for the Seldinger technique. (H) Final custom-built construction. <a href="https://www.jove.com/files/ftp_upload/66684/66684fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Anesthesia and analgesia
<ol>
	<li>Anesthetize the rat with the medication of choice. 
	NOTE: Here, anesthesia and analgesia were established using ketamine 10% (100 mg/mL), xylazine 2% (20 mg/mL), carprofen 50 mg/mL, and isoflurane. A 1:10 dilution of xylazine and carprofen were used as stock solutions for application.
	<ol>
		<li>Induce anesthesia by placing the animal in an induction box flooded with 100% oxygen at an inflow rate of 5 L/min complemented by 4% (v) of isoflurane using a vaporizer.</li>
		<li>After unconsciousness was achieved, remove the animal from the induction box and inject (i.p.) it with 100 mg/kg body weight ketamine and 4 mg/kg body weight xylazine. 
		NOTE: This corresponds to 40 mg of ketamine (0.4 mL of the original solution) and 1.6 mg of xylazine (0.08 mL of the original solution or 0.8 mL of the 1:10 solution) for a 400 mg animal.</li>
		<li>Place the animal back into the isoflurane-flooded induction box until the effect of the i.p. injection sets in.</li>
	</ol>
	</li>
	<li>Guarantee proper anesthetic depth by testing the lack of response of the rat to a firm toe pinch.</li>
	<li>Apply ophthalmic ointment to the eyes.</li>
	<li>Assess and maintain the depth of anesthesia during surgery.
	<ol>
		<li>When narcotic effects seemed to diminish over time (about every hour), inject (s.c) the animal with an additional 50 mg/kg body weight ketamine and 1 mg/kg body weight xylazine. This corresponds to 20 mg of ketamine (0.2 mL of the original solution) and 0.4 mg of xylazine (0.2 mL of the 1:10 solution) for a 400 mg animal.</li>
		<li>Provide additional analgesia before thoracotomy with an s.c. injection of 5 mg/kg body weight carprofen. This corresponds to 2 mg of carprofen (0.04 mL of the original solution or 0.4 mL of the 1:10 solution) for a 400 mg animal.</li>
	</ol>
	</li>
</ol>
3. Surgical preparation (Figure 2)
<ol>
	<li>Saturate the animal circulation with oxygen by offering inhalation of 100% oxygen via a transnasal neonatal mask (Figure 2A).</li>
	<li>After 5 min, switch from the transnasal neonatal mask to the construction of a neonatal ventilation bag with a male plastic syringe Luer-lock tip as described above and offer oxygen in overflow with the syringe tip close to the animal nasal tip (Figure 2B).</li>
	<li>Perform a median thoracocervical skin incision across the desired length and expose the surgical situs using the surgical preparation hooks (Figure 2C,D).</li>
	<li>Advance the preparation by blunt dissection with fine scissors through the cervical fascia. Expose the sternocleidomastoid and infrahyoidal muscle and lateralize the median infrahyoidal muscle to the right after left lateral membrane dissection (Figure 2E-G).</li>
	<li>Perform blunt dissection towards the trachea and tunneling of the trachea using overholt clamps (Figure 2H-I).</li>
	<li>Sling the trachea using a silicone vessel loop and double sling the distal trachea using a polyfilament suture for later endotracheal cannula fixation (Figure 2J-N).</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66684/66684fig02.jpg" /> 
Figure 2: Preparation of the trachea. (A) Inhalation of oxygen via mask. (B) Switch from mask to male Luer-lock syringe for oxygen application. (C) Thoracocervical skin incision. (D) Preparation through the cervical fascia. (E) exposure of the sternocleidomastoid muscle and the infrahyoidal muscle. (F) Lateralization of the median infrahyoidal muscle. (G) Blunt dissection towards the trachea. (H) tunneling of the trachea using overholt clamps. (I) Slinging of the trachea using a silicone vessel loop. (J-N) Caudal double slinging of the trachea using a poly filament suture for later endotracheal cannula fixation. <a href="https://www.jove.com/files/ftp_upload/66684/66684fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Intubation procedure (Figure 3)
<ol>
	<li>Caudally lengthen the trachea using atraumatic pliers (Figure 3A,B).</li>
	<li>Perform a partial incision of the trachea of 180&#176; of circumference using scissors (Figure 3C-F).</li>
	<li>Introduce a Seldinger guide wire with the flexible end into the trachea (Figure 3G-H) and subsequently introduce the modified intravenous catheter of appropriate girth into the trachea guided by the Seldinger wire (Figure 3I-N).</li>
	<li>Remove the Seldinger wire (Figure 3O), connect the Luer-lock tip of the tracheal catheter to the modified ventilation bag, and start gentle lung-protective manual ventilation with high frequency and low tidal volume (Figure 3P). Perform manual ventilation as long as median thoracotomy has not been performed and the lung is not in plain sight.</li>
	<li>Tie the previously placed locking suture with a sliding knot to avoid aerial leakage and accidental catheter removal (Figure 3Q-T).</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66684/66684fig03.jpg" /> 
Figure 3: Endotracheal intubation via tracheotomy. (A) Initial situation after preparation. (B) Caudal lengthening of the trachea using atraumatic pliers. (C-F) Partial incision of the trachea of 180&#176; of circumference. (G,H) Introduction of a Seldinger guide wire into the trachea. (I-M) Introduction of the modified intravenous catheter into the trachea guided by the Seldinger wire. (N-O) Removal of the Seldinger wire. (P) Luer-lock connection of the tracheal catheter to the modified ventilation bag. (Q-T) Tying of the previously placed locking suture to avoid accidental catheter removal. <a href="https://www.jove.com/files/ftp_upload/66684/66684fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Manual ventilation and median thoracotomy (Figure 4)
<ol>
	<li>Start the median thoracotomy from the xiphoid using blunt and stable material scissors (Figure 4A,B) and continue cranially through the sternum. Always pause the ventilation for a few seconds when advancing with the scissors substernally and when cutting in order to avoid trauma to the lung (Figure 4C-E).</li>
	<li>Gain thoracic exposure using surgical preparation hooks and remove mediastinal serosal membranes (Figure 4F-H).</li>
	<li>Continue manual ventilation with desired frequency and tidal volume and expand monitoring (e.g., establishing an intra-arterial blood pressure measurement, installing a central venous or pulmonary arterial catheter depending on the purpose or respective research question) (Figure 4I-L).</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66684/66684fig04.jpg" /> 
Figure 4: Manual ventilation and median thoracotomy. (A) initial situation after intubation. (B-E) Median thoracotomy starting from the xiphoid.(F-I) Thoracic exposure using surgical preparation hooks. (J) Final setup during ventilation. (K-L) the process of manual ventilation. <a href="https://www.jove.com/files/ftp_upload/66684/66684fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Euthanasia
<ol>
	<li>Achieve euthanasia by abrupt sharp cardiectomy using scissors in the deeply anesthetized animal.</li>
</ol>

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

<ol><li>Czigany, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Limb+remote+ischemic+conditioning+of+the+recipient+protects+the+liver+in+a+rat+model+of+arterialized+orthotopic+liver+transplantation%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">Limb remote ischemic conditioning of the recipient protects the liver in a rat model of arterialized orthotopic liver transplantation.</a> PLoS One. 13 (4), e0195507(2018).</li>
<li>Czigány, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Improving+research+practice+in+rat+orthotopic+and+partial+orthotopic+liver+transplantation%3A+A+review%2C+recommendation%2C+and+publication+guide%5BTitle%5D)+AND+%22Eur+Surg+Res%22%5BJournal%5D)">Improving research practice in rat orthotopic and partial orthotopic liver transplantation: A review, recommendation, and publication guide.</a> Eur Surg Res. 55 (1-2), 119-138 (2015).</li>
<li>Nagai, K., Yagi, S., Uemoto, S., Tolba, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Surgical+procedures+for+a+rat+model+of+partial+orthotopic+liver+transplantation+with+hepatic+arterial+reconstruction%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Surgical procedures for a rat model of partial orthotopic liver transplantation with hepatic arterial reconstruction.</a> J Vis Exp. 73, e4376(2013).</li>
<li>Jahshan, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+novel+rat+model+for+assessment+of+laryngotracheal+injury+following+transoral+intubation%5BTitle%5D)+AND+%22Int+J+Pediatr+Otorhinolaryngol%22%5BJournal%5D)">A novel rat model for assessment of laryngotracheal injury following transoral intubation.</a> Int J Pediatr Otorhinolaryngol. 113, 4-10 (2018).</li>
<li>Lamoureux, L., Radhakrishnan, J., Gazmuri, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+rat+model+of+ventricular+fibrillation+and+resuscitation+by+conventional+closed-chest+technique%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">A rat model of ventricular fibrillation and resuscitation by conventional closed-chest technique.</a> J Vis Exp. 98, 52413(2015).</li>
<li>Wang, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Autoinducer-2+of+streptococcus+mitis+as+a+target+molecule+to+inhibit+pathogenic+multi-species+biofilm+formation+in+vitro+and+in+an+endotracheal+intubation+rat+model%5BTitle%5D)+AND+%22Front+Microbiol%22%5BJournal%5D)">Autoinducer-2 of streptococcus mitis as a target molecule to inhibit pathogenic multi-species biofilm formation in vitro and in an endotracheal intubation rat model.</a> Front Microbiol. 7, 88(2016).</li>
<li>Jahshan, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+novel+rat+model+for+tracheal+mucosal+damage+assessment+of+following+long+term+intubation%5BTitle%5D)+AND+%22Int+J+Pediatr+Otorhinolaryngol%22%5BJournal%5D)">A novel rat model for tracheal mucosal damage assessment of following long term intubation.</a> Int J Pediatr Otorhinolaryngol. 128, 109738(2020).</li>
<li>Rivard, A. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rat+intubation+and+ventilation+for+surgical+research%5BTitle%5D)+AND+%22J+Invest+Surg%22%5BJournal%5D)">Rat intubation and ventilation for surgical research.</a> J Invest Surg. 19 (4), 267-274 (2006).</li>
<li>Na, N., Zhao, D. Q., Huang, Z. Y., Hong, L. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+improved+method+for+rat+intubation+and+thymectomy%5BTitle%5D)+AND+%22Chin+Med+J+%28Engl%29%22%5BJournal%5D)">An improved method for rat intubation and thymectomy.</a> Chin Med J (Engl). 124 (17), 2723-2727 (2011).</li>
<li>Cicero, L., Fazzotta, S., Palumbo, V. D., Cassata, G., Lo Monte, A. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Anesthesia+protocols+in+laboratory+animals+used+for+scientific+purposes%5BTitle%5D)+AND+%22Acta+Biomed%22%5BJournal%5D)">Anesthesia protocols in laboratory animals used for scientific purposes.</a> Acta Biomed. 89 (3), 337-342 (2018).</li>
<li>Fuentes, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Videoendoscopic+endotracheal+intubation+in+the+rat%3A+A+comprehensive+rodent+model+of+laparoscopic+surgery%5BTitle%5D)+AND+%22J+Surg+Res%22%5BJournal%5D)">Videoendoscopic endotracheal intubation in the rat: A comprehensive rodent model of laparoscopic surgery.</a> J Surg Res. 122 (2), 240-248 (2004).</li>
<li>Bryda, E. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+mighty+mouse%3A+The+impact+of+rodents+on+advances+in+biomedical+research%5BTitle%5D)+AND+%22Mo+Med%22%5BJournal%5D)">The mighty mouse: The impact of rodents on advances in biomedical research.</a> Mo Med. 110 (3), 207-211 (2013).</li>
<li>Lazopoulos, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Open+thoracotomy+for+pneumothorax%5BTitle%5D)+AND+%22J+Thorac+Dis%22%5BJournal%5D)">Open thoracotomy for pneumothorax.</a> J Thorac Dis. 7 (S1), S50-S55 (2015).</li>
<li>Rendell, V. R., Giamberardino, C., Li, J., Markert, M. L., Brennan, T. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Complete+thymectomy+in+adult+rats+with+non-invasive+endotracheal+intubation%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Complete thymectomy in adult rats with non-invasive endotracheal intubation.</a> J Vis Exp. 94, 52152(2014).</li>
<li>Thomas, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Endotracheal+intubation+in+mice+via+direct+laryngoscopy+using+an+otoscope%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Endotracheal intubation in mice via direct laryngoscopy using an otoscope.</a> J Vis Exp. 86, 50269(2014).</li>
<li>Nelson, A. M., Nolan, K. E., Davis, I. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Repeated+orotracheal+intubation+in+mice%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Repeated orotracheal intubation in mice.</a> J Vis Exp. 157, 60844(2020).</li>
<li>Das, S., Macdonald, K., Chang, H. Y., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+simple+method+of+mouse+lung+intubation%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">A simple method of mouse lung intubation.</a> J Vis Exp. 73, e50318(2013).</li>
<li>Massick, D. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Quantification+of+the+learning+curve+for+percutaneous+dilatational+tracheotomy%5BTitle%5D)+AND+%22Laryngoscope%22%5BJournal%5D)">Quantification of the learning curve for percutaneous dilatational tracheotomy.</a> Laryngoscope. 110 (2 Pt 1), 222-228 (2000).</li>
<li>Giugliano, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Learning+curve+for+translaryngeal+tracheotomy+in+head+and+neck+surgery%5BTitle%5D)+AND+%22Laryngoscope%22%5BJournal%5D)">Learning curve for translaryngeal tracheotomy in head and neck surgery.</a> Laryngoscope. 111 (4 Pt 1), 628-633 (2001).</li>
<li>Mort, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Emergency+tracheal+intubation%3A+Complications+associated+with+repeated+laryngoscopic+attempts%5BTitle%5D)+AND+%22Anesth+Analg%22%5BJournal%5D)">Emergency tracheal intubation: Complications associated with repeated laryngoscopic attempts.</a> Anesth Analg. 99 (2), 607-613 (2004).</li>
<li>Hasegawa, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Association+between+repeated+intubation+attempts+and+adverse+events+in+emergency+departments%3A+An+analysis+of+a+multicenter+prospective+observational+study%5BTitle%5D)+AND+%22Ann+Emerg+Med%22%5BJournal%5D)">Association between repeated intubation attempts and adverse events in emergency departments: An analysis of a multicenter prospective observational study.</a> Ann Emerg Med. 60 (6), 749-754.e742 (2012).</li>
<li>Titu, I. M., Delaca, G. B., Teterea, F., Ciulic, S. A., Palade, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Percutaneous+tracheostomy+using+the+Seldinger+technique%5BTitle%5D)+AND+%22Multimed+Man+Cardiothorac+Surg%22%5BJournal%5D)">Percutaneous tracheostomy using the Seldinger technique.</a> Multimed Man Cardiothorac Surg. 2023, (2023).</li>
<li>Garry, B. P., Bivens, H. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+Seldinger+technique%5BTitle%5D)+AND+%22J+Cardiothorac+Anesth%22%5BJournal%5D)">The Seldinger technique.</a> J Cardiothorac Anesth. 2 (3), 403(1988).</li>
<li>Carbone, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Estimating+mouse+and+rat+use+in+American+laboratories+by+extrapolation+from+animal+welfare+act-regulated+species%5BTitle%5D)+AND+%22Sci+Rep%22%5BJournal%5D)">Estimating mouse and rat use in American laboratories by extrapolation from animal welfare act-regulated species.</a> Sci Rep. 11 (1), 493(2021).</li>
<li>Su, C. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Efficacious+and+safe+orotracheal+intubation+for+laboratory+mice+using+slim+torqueable+guidewire-based+technique%3A+Comparisons+between+a+modified+and+a+conventional+method%5BTitle%5D)+AND+%22BMC+Anesthesiol%22%5BJournal%5D)">Efficacious and safe orotracheal intubation for laboratory mice using slim torqueable guidewire-based technique: Comparisons between a modified and a conventional method.</a> BMC Anesthesiol. 16, 5(2016).</li>
<li>Clary, E. M., O'halloran, E. K., De La Fuente, S. G., Eubanks, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Videoendoscopic+endotracheal+intubation+of+the+rat%5BTitle%5D)+AND+%22Lab+Anim%22%5BJournal%5D)">Videoendoscopic endotracheal intubation of the rat.</a> Lab Anim. 38 (2), 158-161 (2004).</li>
<li>Ghali, M. G. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microsurgical+technique+for+tracheostomy+in+the+rat%5BTitle%5D)+AND+%22MethodsX%22%5BJournal%5D)">Microsurgical technique for tracheostomy in the rat.</a> MethodsX. 5, 61-67 (2018).</li>
</ol>

The authors have nothing to disclose.

In the US alone, about 110 million rats and mice are used in animal experiments per year24. While no reliable statistics can be found, a relevant percentage of these will receive supported ventilation, especially in the fields of pulmonary and cardiac research as well as anesthesiology. Most of these animals receive non-invasive endoorotracheal intubation, which entails a number of challenges. These include slower learning curves, higher rates of intubation failure, greater numbers of reintubations, and associated respiratory complications as well as respiratory leakages25. Non-invasive intubation will, therefore, generally take more experienced research staff, more advanced technology such as fiber optics, capnometry, and mechanical ventilators, and greater financial and materialistic resources26. The whole aspect of intubation is, therefore, a meaningful confounder relevant in subsequent statistical analysis and interindividual heterogeneity of results. However, non-invasive endoorotracheal intubation is often the only feasible option in longitudinal experiment designs. 
Alternatives are tracheostomy for required long-term tracheal access in longitudinal experiments or surgical tracheotomy for "single-use" in non-survival experiments. While protocols on the microsurgical technique for tracheostomy in rats are available27, there are currently no protocols for endotracheal intubation via tracheotomy in non-survival rat models.
The herein-described standardized procedure for endotracheal intubation via tracheotomy and subsequent thoracotomy in rats, therefore, represents a valuable method for researchers engaged in non-survival studies. This procedure requires neither expert personnel nor extensive training. It has an intubation success rate that is much higher than endoorotracheal intubation and allows for much more controlled procedural steps. 
Limitations of the presented techniques mainly include the invasiveness of the procedure and, consecutively, the exclusive compatibility with non-survival experiments. Due to the large defects in skin and soft tissue, as well as the incisional tracheal fistula, extubation and recovery from this procedure are neither feasible nor recommended, especially when combined with thoracotomy.
When troubleshooting common challenges encountered during the procedure, we would like to draw attention to the following points and recommendations: one should prepare equipment and medication as extensively as possible. The experimenter should perform hemostatic control meticulously by fine preparation and using non-traumatic cotton swabs. The experimenter should prepare the trachea over a length of at least 0.5 cm in order to allow for sufficient mobility and use the silicone vessel loop in order to not lose the trachea after incision, as this can result in aspiration and suffocation when the trachea is lost in the tissue depth due to backdrop phenomenon. Only partially incise the trachea with one clear stroke of the scissors. Ideally, one should incise 180&#176; of circumference. Below this might result in failure to intubate. Above this might result in snapping of the trachea with retraction of both ends into the depth of the tissue and subsequent suffocation. When tying the polyfilament suture to secure the intubation tube, a sliding knot should be used in order to ensure sufficient tightness to avoid air leakage or accidental tube dislocation while at the same time avoiding unintended closure of the tube. Leaving the Seldinger wire in place during suturing can avoid unintended closure. When performing manual ventilation using the ventilation bag, one should strictly avoid barotrauma of the lung by high-tidal volumes. Instead, one should rather aim for lung-protective, high-frequent, and low-tidal ventilation, especially as long as thoracotomy has not been performed and the lung volumes cannot be evaluated visually.
By offering a detailed and reproducible methodology, this protocol facilitates the standardization of experimental procedures, improving data reliability and comparability across studies. As a result, this method contributes to the refinement of scientific techniques in preclinical research, ultimately enhancing the translational relevance of findings in the fields of physiology, pharmacology, surgical and biomedical sciences.

In general, rodents are able to compensate for respiratory distress for much longer than patients would. They can remain cardiocirculatory stable and sufficient under spontaneous ventilation for much longer and during more invasive procedures, e.g., even through liver transplantation, which is known to be one of the procedures most stressful for cardiopulmonary circulation1,2,3.
However, endotracheal intubation is a basic requirement for translational research in rat models for various settings and interventions originating from a variety of biomedical fields and is essential in today&#39;s scientific landscape4,5,6,7,8,9,10,11. While the majority of experimental laboratory animal work performed in rats in general still does not require invasive ventilation12, there are certain experimental setups that require intubation and controlled ventilatory assistance. These experimental setups include any experiment requiring a secured airway, high ventilation pressures, and laparoscopy or direct access to the thoracic cavity and organs.
Especially in the case of required access to the thoracic cavity and organs, endotracheal intubation is imperative as the collapse of the lung will result in a fatal respiratory insufficiency once the negative pressure in the intrapleural space and the inspiration mechanism by the intercostal muscles and the diaphragm are lost13.
While there are many publications on methods for non-invasive endoorotracheal intubation in rodents14,15,16,17, there seems to be a lack of procedure protocols for invasive endotracheal intubation by tracheotomy. Despite the invasiveness of the latter, which only allows for application in non-survival animal models, there are great advantages of intubation via tracheotomy. These include a steeper learning curve, higher and more persistent success rates, less required technical equipment, and improved success monitoring opportunities18,19.
Intubation success is essential for outcome. While various false intubations, as well as repeated intubation attempts in patients, are clearly associated with adverse events and complications as serious as death20,21, such events are also deleterious in test animals. In the best case, they represent a strong confounder variable in the experiment, but they can also lead to the unnecessary loss of the animal. Therefore, it makes sense to increase intubation success rates at the cost of non-invasiveness if the experimental setup and strategy allow it.
This standardized protocol of intubation via surgical tracheotomy offers several advantages, including reduced variability in intubation success rates, minimized effects on the cardiorespiratory physiology parameters, and full investigational control and manipulation of parameters. It aids in providing a secured airway, especially for inexperienced researchers in the procedure of endotracheal intubation via direct laryngoscopy, and allows researchers to conduct non-survival studies with highly controlled conditions. We highlight key anatomical landmarks and provide insights into troubleshooting common challenges encountered during the procedure.

<table><tbody><tr><td>Ambu Spur II Single-Use ventilation bag for neonates</td><td>Meier Medizintechnik</td><td>335 102 000</td><td>Ventilation bag</td></tr><tr><td>BD Microlance 3 cannula 20 G</td><td>BD (Beckton, Dickinson)</td><td>301300</td><td>Cannula</td></tr><tr><td>BD Microlance Discardit II 20 mL Syringe</td><td>BD (Beckton, Dickinson)</td><td>300296</td><td>Plastic syringe</td></tr><tr><td>Fixation rods</td><td>Legefirm</td><td>&amp;lrm;500343896</td><td>Tuning forks used as y-shaped metal fixation rods</td></tr><tr><td>Heat-shrinking tube</td><td>Sekesoer</td><td>RSG-400</td><td>Heat-shrinking tube</td></tr><tr><td>Intravenous catheter</td><td>BD (Beckton, Dickinson)</td><td>393230</td><td>BD Venflon Pro Safety intravenous catheter 14 G; shortened using scissors; alternatively 16 G or 18 G can be used</td></tr><tr><td>Plastic perfusor tube</td><td>M. Schilling GmbH</td><td>S702NC150</td><td>Connecting tube COEX 150 cm</td></tr><tr><td>Polyfilament suture</td><td>COVIDIEN</td><td>CL331</td><td>Polyfilament surgical suture recommended with a strength of 1 to 2; needle can be removed</td></tr><tr><td>Royal Gardineer Heating Pad Size S, 20 Watt</td><td>Royal Gardineer</td><td>IP67</td><td>Heating pad</td></tr><tr><td>Seldinger guide wire</td><td>VYGON</td><td>115.798</td><td>Metal guide wire from the arterial leadercath set</td></tr><tr><td>Silicone vessel loop tie</td><td>SERAG WIESSNER</td><td>SL26</td><td>Silicone vessel loop tie 2.5 mm red</td></tr><tr><td>Spraque Dawley rats</td><td>Janvier Labs</td><td></td><td>Male rats weighing 400 grams</td></tr><tr><td>Steel plate</td><td>Maschinenbau Feld GmbH</td><td>C010206</td><td>Galvanized sheet plate, 40 x 50 cm, thickness 4.0 mm</td></tr></tbody></table>

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.

Endotracheal intubation via tracheotomy and subsequent thoracotomy was performed in 10 male rats (mean weight 405 &#177; 30 g) for non-survival applications. The procedure was performed by a second-year surgical resident. The success rate defined by survival over 20 min of intubation and ventilation was 100%. The mean duration of the preparation and intubation procedure from skin incision until fixation of the intubation tube was 6:55 &#177; 0:53 min (Table 1).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Number of animals used</td>
			<td>10</td>
		</tr>
		<tr>
			<td>Body weight (mean and standard deviation)</td>
			<td>405 &#177; 30 g</td>
		</tr>
		<tr>
			<td>Intubation success rate</td>
			<td>100%</td>
		</tr>
		<tr>
			<td>Duration of procedure (mean and standard deviation)</td>
			<td>6:55 &#177; 0:53 min</td>
		</tr>
		<tr>
			<td>Mean oxygen saturation</td>
			<td>96%</td>
		</tr>
		<tr>
			<td>Minimum oxygen saturation</td>
			<td>92%</td>
		</tr>
	</tbody>
</table>
Table 1: Representative results. Representative results of the protocol in 10 animals.
Saturation was monitored through a single-use self-stick pulse-oximetry probe on the hind leg and never fell below 92%. Spontaneous ventilation was contained until the incision of the diaphragm. All rats survived the 20 min required for the experimental measurements until the intentional euthanasia through abrupt sharp cardiectomy (Figure 5).
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66684/66684fig05.jpg" /> 
Figure 5: Peripheral oxygen saturation. Peripheral oxygen saturation measured over the whole duration of this procedure. <a href="https://www.jove.com/files/ftp_upload/66684/66684fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Read this Scientific Article Protocol about Author Spotlight: Hyperspectral Imaging for Spectral Analysis of Thoracic Organs in Clinical and Animal Models at JoVE.com

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

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 ...

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Research

JoVE Journal

Medicine

555 Views.  Heidelberg University Hospital. 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.

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

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 ...

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.

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 ...

Immunology and Infection

Induction and Assessment of Ischemia-reperfusion Injury in Langendorff-perfused Rat Hearts

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for myocardial infarction and cardiac complications of thoracic surgery. The ability of certain drugs to precondition the myocardium against ischemia reperfusion injury has led to multiple clinical trials, with little success. The isolated heart model allows acute observation of the functional effects of ischemia reperfusion injury in real time, including the effects of various pharmacological interventions administered at any time-point before or within the ischemia-reperfusion injury window. Since brief periods of ischemia can precondition the heart against ischemic injury, in situ aortic cannulation is performed to allow for functional assessment of non-preconditioned myocardium. A saline filled balloon is placed into the left ventricle to allow for real-time measurement of pressure generation. Ischemic injury is simulated by the cessation of perfusion buffer flow, followed by reperfusion. The duration of both ischemia and reperfusion can be modulated to examine biochemical events at any given time-point. Although the Langendorff isolated heart model does not allow for the consideration of systemic events affecting ischemia and reperfusion, it is an excellent model for the examination of acute functional and biochemical events within the window of ischemia reperfusion injury as well as the effect of pharmacological intervention on cardiac pre- and postconditioning. The goal of this protocol is to demonstrate how to perform in situ aortic cannulation and heart excision followed by ischemia/reperfusion injury in the Langendorff model.

The isolated rat heart is an enduring model for ischemia reperfusion injury. Here, we describe the process of harvesting the beating heart from a rat via in situ aortic cannulation, Langendorff perfusion of the heart, simulated ischemia-reperfusion injury, and infarct staining to confirm the extent of ischemic insult.

The biochemical events surrounding ischemia reperfusion injury in the acute setting are of great importance to furthering novel treatment options for ...

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 ...

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute respiratory distress syndrome (ARDS), characterized by neutrophilic alveolitis, and injury to both alveolar epithelium and vascular endothelium. Clinically, ARDS is defined by acute onset of hypoxemia, bilateral patchy pulmonary infiltrates and non-cardiogenic pulmonary edema. Human studies have provided us with valuable information about the physiological and inflammatory changes in the lung caused by ARDS, which has led to various hypotheses about the underling mechanisms. Unfortunately, difficulties determining the etiology of ARDS, as well as a wide range of pathophysiology have resulted in a lack of critical information that could be useful in developing therapeutic strategies.
Translational animal models are valuable when their pathogenesis and pathophysiology accurately reproduce a concept proven in both in vitro and clinical settings. Although large animal models (e.g., sheep) share characteristics of the anatomy of human trachea-bronchial tree, murine models provide a host of other advantages including: low cost; short reproductive cycle lending itself to greater data acquisition; a well understood immunologic system; and a well characterized genome leading to the availability of a variety of gene deletion and transgenic strains. A robust model of low pH induced ARDS requires a murine ALI that targets mainly the alveolar epithelium, secondarily the vascular endothelium, as well as the small airways leading to the alveoli. Furthermore, a reproducible injury with wide differences between different injurious and non-injurious insults is important.
The murine gastric acid aspiration model presented here using hydrochloric acid employs an open tracheostomy and recreates a pathogenic scenario that reproduces the low pH pneumonitis injury in humans. Additionally, this model can be used to examine interaction of a low pH insult with other pulmonary injurious entities (e.g., food particles, pathogenic bacteria).

This protocol induces acute lung injury in a mouse that has close fidelity to the pathogenesis of acid pneumonitis observed in humans. We generate a maximal acute nonlethal low pH lung injury and account for differences in rodent-human anatomic respiratory structure using an open tracheostomy coupled with circumferential pressure release.

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute ...

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 ...

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 ...

A Minimally Invasive Method for Intratracheal Instillation of Drugs in Neonatal Rodents to Treat Lung Disease

Treatment of neonatal rodent with drugs instilled directly into the trachea could serve as a valuable tool to study the impact of a locally administered drug. This has direct translational impact because surfactant and drugs are administered locally into the lungs. Though the literature has many publications describing minimally invasive transoral intubation of adult mice and rats in therapeutic experiments, this approach in neonatal rat pups is lacking. The small size of orotracheal region/pharynx in the pups makes visualization of laryngeal lumen (vocal cords) difficult, contributing to the variable success rate of intratracheal drug delivery. We hereby demonstrate effective oral intubation of neonatal rat pup - a technique that is non-traumatic and minimally-invasive, so that it can be used for serial administration of drugs. We used an operating otoscope with an illumination system and a magnifying lens to visualize the tracheal opening of the rat neonates. The drug is then instilled using a 1 mL syringe connected to a pipette tip. The accuracy of the delivery method was demonstrated using Evans blue dye administration. This method is easy to get trained in and could serve as an effective way to instill drugs into trachea. This method could also be used for administration of inoculum or agents to simulate disease conditions in animals and, also, for cell-based treatment strategies for various lung diseases.

This technique of instilling drugs directly into the trachea of neonatal rodents is important in studying the impact of locally administered drugs or biologicals on neonatal lung diseases. Additionally, this method can also be used for inducing lung injury in animal models.

Treatment of neonatal rodent with drugs instilled directly into the trachea could serve as a valuable tool to study the impact of a locally administered ...

Endotracheal Intubation of Rabbits Using a Polypropylene Guide Catheter

Endotracheal intubation in rabbits can be challenging due to their unusual anatomy. Achieving a patent airway during anesthesia is critical for the avoidance of airway obstruction, prevention of gastric tympany, and to allow ventilatory support. Due to the difficulty of intubation, alternative methods such as the use of laryngeal mask airways or laryngeal tubes have been explored. However, these methods do not result in direct access to the trachea and thus may present a risk for development of complications. In addition, lack of direct intubation of the trachea can result in personnel exposure to waste anesthetic gases. Numerous methods for endotracheal intubation have been described, including blind placement, use of a fiberoptic laryngoscope or endoscope, and cricoid placement. Despite these numerous publications, many still struggle to achieve success. Here we provide a detailed description of an intubation technique that can be taught with minimal training with a short time to proficiency. Briefly, after administration of injectable anesthesia and proper positioning of the rabbit, a polypropylene catheter is placed into the trachea by direct visualization using a laryngoscope. The catheter is then used as a guide to direct the endotracheal tube into the trachea. This method allows for intubation without the need for expensive equipment and can be performed by a single individual without the need for an assistant. In conclusion, this technique can be easily taught and performed at very little cost in any clinical or research setting.

Endotracheal intubation in rabbits is challenging due to their unusual anatomy. Here we present a technique for direct intubation of the trachea using a polypropylene catheter as a guide. This method utilizes relatively inexpensive supplies, requires minimal training, and can be easily performed in any clinical setting.

Endotracheal intubation in rabbits can be challenging due to their unusual anatomy. Achieving a patent airway during anesthesia is critical for the ...

Biology

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.

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

Summary

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Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

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An Improved Method for Rapid Intubation of the Trachea in Mice

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

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

Repeated Orotracheal Intubation in Mice

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