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

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>&#8206;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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422 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

A Novel Rescue Technique for Difficult Intubation and Difficult Ventilation

We describe a novel non surgical technique to maintain oxygenation and ventilation in a case of difficult intubation and difficult ventilation, which works especially well with poor mask fit.
Can not intubate, can not ventilate" (CICV) is a potentially life threatening situation. In this video we present a simulation of the technique we used in a case of CICV where oxygenation and ventilation were maintained by inserting an endotracheal tube (ETT) nasally down to the level of the naso-pharynx while sealing the mouth and nares for successful positive pressure ventilation.
A 13 year old patient was taken to the operating room for incision and drainage of a neck abcess and direct laryngobronchoscopy. After preoxygenation, anesthesia was induced intravenously. Mask ventilation was found to be extremely difficult because of the swelling of the soft tissue. The face mask could not fit properly on the face due to significant facial swelling as well. A direct laryngoscopy was attempted with no visualization of the larynx. Oxygen saturation was difficult to maintain, with saturations falling to 80%. In order to oxygenate and ventilate the patient, an endotracheal tube was then inserted nasally after nasal spray with nasal decongestant and lubricant. The tube was pushed gently and blindly into the hypopharynx. The mouth and nose of the patient were sealed by hand and positive pressure ventilation was possible with 100% O2 with good oxygen saturation during that period of time. Once the patient was stable and well sedated, a rigid bronchoscope was introduced by the otolaryngologist showing extensive subglottic and epiglottic edema, and a mass effect from the abscess, contributing to the airway compromise. The airway was secured with an ETT tube by the otolaryngologist.This video will show a simulation of the technique on a patient undergoing general anesthesia for dental restorations.

We describe a technique to maintain oxygenation and ventilation using an endotracheal tube inserted nasally to the level of the naso-pharynx while sealing the mouth and nares for successful positive pressure ventilation.

We describe a novel non surgical technique to maintain oxygenation and ventilation in a case of difficult intubation and difficult ventilation, which ...

Guidelines for Elective Pediatric Fiberoptic Intubation

Fiberoptic intubation in pediatric patients is often required especially in difficult airways of syndromic patients i.e. Pierre Robin Syndrome. Small babies will desaturate very quickly if ventilation is interrupted mainly to high metabolic rate. We describe guidelines to perform a safe fiberoptic intubation while maintaining spontaneous breathing throughout the procedure. Steps requiring the use of propofol pump, fentanyl, glycopyrrolate, red rubber catheter, metal insuflation hook, afrin, lubricant and lidocaine spray are shown.

We describe guidelines to perform a safe and efficient elective fiberoptic intubation in pediatric patients while maintaining spontaneous ventilation.

Fiberoptic intubation in pediatric patients is often required especially in difficult airways of syndromic patients i.e. Pierre Robin Syndrome. Small ...

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 ...

A Simple Method of Mouse Lung Intubation

A simple procedure to intubate mice for pulmonary function measurements would have several advantages in longitudinal studies with limited numbers or expensive animal. One of the reasons that this is not done more routinely is that it is relatively difficult, despite there being several published studies that describe ways to achieve it. In this paper we demonstrate a procedure that eliminates one of the major hurdles associated with this intubation, that of visualizing the trachea during the entire time of intubation. The approach uses a 0.5 mm fiberoptic light source that serves as an introducer to direct the intubation cannula into the mouse trachea. We show that it is possible to use this procedure to measure lung mechanics in individual mice over a time course of at least several weeks. The technique can be set up with relatively little expense and expertise, and it can be routinely accomplished with relatively little training. This should make it possible for any laboratory to routinely carry out this intubation, thereby allowing longitudinal studies in individual mice, thereby minimizing the number of mice needed and increasing the statistical power by using each mouse as its own control.

This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

A  simple procedure to intubate mice for pulmonary function measurements would  have several advantages in longitudinal studies with limited numbers or  ...

Collecting Saliva and Measuring Salivary Cortisol and Alpha-amylase in Frail Community Residing Older Adults via Family Caregivers

Salivary measures have emerged in bio-behavioral research that are easy-to-collect, minimally invasive, and relatively inexpensive biologic markers of stress. This article we present the steps for collection and analysis of two salivary assays in research with frail, community residing older adults-salivary cortisol and salivary alpha amylase. The field of salivary bioscience is rapidly advancing and the purpose of this presentation is to provide an update on the developments for investigators interested in integrating these measures into research on aging. Strategies are presented for instructing family caregivers in collecting saliva in the home, and for conducting laboratory analyses of salivary analytes that have demonstrated feasibility, high compliance, and yield quality specimens. The protocol for sample collection includes: (1) consistent use of collection materials; (2) standardized methods that promote adherence and minimize subject burden; and (3) procedures for controlling certain confounding agents. We also provide strategies for laboratory analyses include: (1) saliva handling and processing; (2) salivary cortisol and salivary alpha amylase assay procedures; and (3) analytic considerations.

We demonstrate: (1) procedures for collection of salivary samples in cognitive impaired older adults by family caregivers in the home setting, (2) procedures for measuring stress activity via salivary cortisol and alpha amylase, and (3) representative profiles. Protocols that allow researchers to study stress-linked processes advance our understanding of biological sensitivity and susceptibility.

Salivary measures have emerged in bio-behavioral research  that are easy-to-collect, minimally invasive, and relatively inexpensive  biologic markers of ...

Intubation-mediated Intratracheal (IMIT) Instillation: A Noninvasive, Lung-specific Delivery System

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences between the murine and human respiratory systems, especially in their upper respiratory tracts (URT). In some models, these differences in the murine nasal cavity can have a significant impact on disease progression and presentation in the lower respiratory tract (LRT) when using intranasal instillation techniques, potentially limiting the usefulness of the mouse model to study these diseases. For these reasons, it would be advantageous to develop a technique to instill bacteria directly into the mouse lungs in order to study LRT disease in the absence of involvement of the URT. We have termed this lung specific delivery technique intubation-mediated intratracheal (IMIT) instillation. This noninvasive technique minimizes the potential for instillation into the bloodstream, which can occur during more invasive traditional surgical intratracheal infection approaches, and limits the possibility of incidental digestive tract delivery. IMIT is a two-step process in which mice are first intubated, with an intermediate step to ensure correct catheter placement into the trachea, followed by insertion of a blunt needle into the catheter to mediate direct delivery of bacteria into the lung. This approach facilitates a &#62;98% efficacy of delivery into the lungs with excellent distribution of reagent throughout the lung. Thus, IMIT represents a novel approach to study LRT disease and therapeutic delivery directly into the lung, improving upon the ability to use mice as surrogates to study human respiratory disease. Furthermore, the accuracy and reproducibility of this delivery system also makes it amenable to Good Laboratory Practice Standards (GLPS), as well as delivery of a wide range of reagents which require high efficiency delivery to the lung.

Intubation-mediated Intratracheal IMIT Instillation: A Noninvasive, Lung-specific Delivery System

Intubation-mediated intratracheal (IMIT) instillation of reagents is an excellent, noninvasive method for studying respiratory disease, as well as a method for instilling therapeutic reagents directly into the lung. It is a rapid and highly reproducible method which is suitable for preclinical testing.

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences ...

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 ...

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 ...

Blind Endotracheal Intubation in Neonatal Rabbits

The newborn rabbit is a useful animal model for various pathologies and procedures. Airway management of the rabbit is complex due to its anatomical characteristics, which is further complicated in the case of the newborn. Of the different methods of advanced airway management, endotracheal intubation is less aggressive than tracheostomy, and is more feasible than supraglottic management given the lack of supraglottic devices of such a small size. As direct glottis visualization is very difficult in animals this size, this blind intubation model is presented as an effective alternative, especially for experiments requiring prolonged anesthesia. Using this method, we performed blind intubations with a 90% success rate.

We describe a technique of endotracheal intubation in newborn rabbits after esophageal catheterization with a gastric tube.

The newborn rabbit is a useful animal model for various pathologies and procedures. Airway management of the rabbit is complex due to its anatomical ...

Endotracheal Intubation Using a Flexible Intubation Endoscope as a Standardized Model for Safe Airway Management in Swine

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway or high ventilation pressures. Endotracheal intubation is a challenging skill,&#160;requiring a minimum number of successful endotracheal intubations to achieve a high success rate under optimal conditions, which is often unachievable for non-anaesthesiology researchers. Due to the specific porcine airway anatomy, a difficult airway can usually be assumed. The impossibility of establishing a secure airway can result in injury, adverse events, or death of the laboratory animal. Using a prospective, randomized, controlled evaluation approach, it has been shown that fiberoptic-assisted endotracheal intubation takes longer but has a higher first-pass success rate than conventional intubation without causing clinically relevant drops in oxygen saturation. This model presents a standardized regimen for endoscopically guided endotracheal intubation, providing a secured airway, especially for researchers who are inexperienced in the technique of endotracheal intubation via direct laryngoscopy. This procedure is expected to minimize animal suffering and unnecessary animal losses.

The use of pigs in research has increased in recent years. Nevertheless, pigs are characterized by difficult airway anatomy. By demonstrating how to perform endoscopically guided endotracheal intubation, the present protocol aims to further increase laboratory animals' safety to avoid animal suffering and unnecessary death.

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway ...