The authors wish to acknowledge those employees and research personnel who have participated in the Rabbit Handling and Anesthesia Training sessions for their feedback while learning this intubation technique.

The intubation procedure outlined below was approved by the Mayo Clinic Institutional Animal Care and Use Committee.
1. Preparation (Figure 1)
<ol>
	<li>Obtain a 5 French polypropylene catheter and cut the non-rounded end so the total length is 8-10 inches. Mark the cut end of the tubing to clearly indicate which end should not be inserted into the trachea (arrow, Figure 1).</li>
	<li>Place a size 0 or 1 Miller laryngoscope blade on the laryngoscope and verify that the light is functioning.</li>
	<li>Draw 0.25 mL of 2% lidocaine into a 1 mL syringe.</li>
	<li>Apply sterile lubricant to the end of an appropriately sized uncuffed endotracheal tube (typically 2.5-4.0 mm ID).</li>
	<li>Prepare supplies for securing the endotracheal tube in place once it is inserted (e.g., umbilical tape).</li>
	<li>Obtain a device to detect air flow at the end of the procedure to confirm successful intubation (#5, Figure 1). This is Optional.</li>
</ol>
2. Anesthesia
<ol>
	<li>Administer injectable anesthesia, such as 35 mg/kg ketamine and 5 mg/kg xylazine intramuscularly using a 3 mL syringe with a 25 gauge needle in the quadriceps muscle of the hind leg of an approximately 4 month old, 3.0-3.5 kg, New Zealand White rabbit. 
	NOTE: This anesthetic regimen is a commonly used injectable combination in young, healthy, specific pathogen-free research rabbits, but based upon clinical condition and/or experimental objectives, a different protocol may be more appropriate.</li>
	<li>After the rabbit is anesthetized, apply sterile ophthalmic lubricant to the eyes to preserve the corneal tissue.</li>
</ol>
3. Endotracheal Intubation
<ol>
	<li>Measure the endotracheal tube from the incisors to the thoracic inlet to determine the proper length for insertion of the endotracheal tube. Placement of the end of the tube at the level of the thoracic inlet assures that the tube opening is past the larynx but cranial to the tracheal bifurcation.</li>
	<li>Position the rabbit sternally on the preparation table with its head slightly extended over the edge of the table (Figure 2). The rabbit should be positioned square with the head and spine in straight alignment.</li>
	<li>Lifting the head up, use a gauze pad to pull the rabbit&#39;s tongue lateral to the rabbit&#39;s right lower incisors (in the diastema), taking care to avoid trauma from the incisors.</li>
	<li>Grasp the laryngoscope and blade with the right hand and place it into the mouth behind the incisors, entering from the rabbit&#39;s left.</li>
	<li>Follow the roof of the mouth with the tip of the blade in a caudal direction until the soft palate is visible.</li>
	<li>With the right hand, tip the laryngoscope forward (bend the wrist, holding the laryngoscope downward) while extending the rabbit&#39;s head back and the neck forward to maintain an open airway and view the larynx (Figure 3A). As obligate nasal breathers, the epiglottis is often above the soft palate, and use of this method compresses the base of the tongue to displace the epiglottis and expose the glottis. To prevent oxygen desaturation, it is important to keep the neck extended as described to maintain an open airway during intubation.</li>
	<li>Release the tongue and switch the laryngoscope blade to the left hand to free up the right hand for placement of the guide catheter.</li>
	<li>While holding the laryngoscope and maintaining view of the glottis (Figure 3B), advance the rounded tip of the guide catheter through the opening past the larynx (Figure 4). The catheter will come to a natural stop at the tracheal bifurcation.</li>
	<li>While maintaining hold of the guide catheter, grasp the endotracheal tube and thread it over the catheter, taking care not to apply force to the catheter by catching it on the endotracheal tube connector. Advance the tube until there is resistance from the vocal cords (Figure 5).</li>
	<li>Instill 0.25 mL of 2% lidocaine into the opening of the endotracheal tube. The lidocaine will travel down the endotracheal tube to locally anesthetize the vocal cords, preventing laryngospasm22 (Figure 6).</li>
	<li>Allow approximately two minutes for the topical anesthesia to take effect, and proceed by gently twisting the endotracheal tube, advancing it through the vocal cords (Figure 7). Once the tube has passed the vocal chords, remove the laryngoscope blade and catheter, taking care not to dislodge the tube.</li>
	<li>Verify placement of the endotracheal tube by observing airflow. There are multiple ways to accomplish this, including placement of a tissue or fur at the end of the tube to detect airflow, looking for fogging of the tube or fogging of a glass/mirror at the end of the tube, and listening for airflow. Devices designed to detect airflow23 or respirations may also be used (Figure 8) to determine whether intubation was successful (see Materials Table for options). Secure the endotracheal tube using a preferred method, such as umbilical tape tied around the tube and head. Attach the anesthesia circuit and auscult both sides of the thorax while giving breaths to assure ventilation to both sides of the lung.</li>
</ol>

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M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laryngeal+mask,+laryngeal+tube,+and+Frova+introducer+in+simulated+difficult+airway.">Laryngeal mask, laryngeal tube, and Frova introducer in simulated difficult airway.</a> J Emerg Med. 48 (2), 254-259 (2015).</li><li>Smith, J. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endotracheal+tubes+versus+laryngeal+mask+airways+in+rabbit+inhalation+anesthesia:+ease+of+use+and+waste+gas+emissions.">Endotracheal tubes versus laryngeal mask airways in rabbit inhalation anesthesia: ease of use and waste gas emissions.</a> Contemp Top Lab Anim Sci. 43 (4), 22-25 (2004).</li><li>Kazakos, G. 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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Teaching+a+new+method+of+rabbit+intubation.">Teaching a new method of rabbit intubation.</a> J Am Assoc Lab Anim Sci. 46 (3), 32-36 (2007).</li><li>Inglis, S., Strunk, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rabbit+anesthesia.">Rabbit anesthesia.</a> Lab Anim (NY). 38 (3), 84-85 (2009).</li><li>Falcao, S. C., Pereira Junior, J. R., Coelho, A. 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O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthesia+of+six-week-old+New+Zealand+White+rabbits+for+thoracotomy.">Anesthesia of six-week-old New Zealand White rabbits for thoracotomy.</a> Contemp Top Lab Anim Sci. 39 (3), 19-22 (2000).</li><li>Stephens DeValle, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Successful+Management+of+Rabbit+Anesthesia+Through+the+Use+of+Nasotracheal+Intubation.">Successful Management of Rabbit Anesthesia Through the Use of Nasotracheal Intubation.</a> Journal of the American Association for Laboratory Animal Science. 48, 166-170 (2009).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> A Guidance on the Use of Topical Anesthetics for Naso/Oropharyngeal and Laryngotracheal Procedures. , (2006).</li><li>Cook, R. T., Stene, J. 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Q., DiNunizio, G., Brodsky, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+study+of+the+tracheotomy+and+endotracheal+intubation+on+the+rabbit+tracheobronchial+tree.">Quantitative study of the tracheotomy and endotracheal intubation on the rabbit tracheobronchial tree.</a> Ann Otol Rhinol. , (1990).</li><li>Kumar, R. A., Boyer, M. I., Bowen, C. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+reliable+method+of+anesthesia+for+extensive+surgery+in+small+rabbits.">A reliable method of anesthesia for extensive surgery in small rabbits.</a> Lab Anim Sci. 43 (3), 265-266 (1993).</li></ol>

The authors have nothing to disclose.

Here we provide a detailed description of a rabbit intubation technique that can be easily mastered in any clinical setting. This technique is similar to previously published direct visualization techniques10,15,20,25 with some additional refinements, including use of a 3.5 mm uncuffed endotracheal tube, and a 5 French urinary catheter as our chosen guide, which has been cut and marked with a permanent marker at the cut end. The rabbit trachea is easily prone to injury and thus the use of uncuffed endotracheal tubes is generally recommended when intubating rabbits to prevent damage to the tracheal mucosa from the pressure of the cuff.26 The 5 French urinary catheter we chose as a guide is small enough to easily pass through the vocal folds without obstructing the view during direct visualization of the larynx. While small, this catheter is stiff enough to remain in place as a guide for the endotracheal tube. The distal tip of the catheter is rounded to help reduce trauma when the bifurcation is reached. The proximal end of the catheter is cut so that the catheter is at a workable length (not excessively long) and to remove the end that widens for attachment to a syringe. The cut end is marked with permanent marker as a visual indicator to not insert this end into the trachea, which may cause damage to the tracheal mucosa.
The primary limitation of this technique is that, because it does require the use of a laryngoscope blade, it may not be feasible for very young rabbits or very small breeds due to smaller mouth size. In addition, some facilities may not have this size of laryngoscope blade readily available. At the author&#39;s institution, this method is used for New Zealand White (&#8805;6 weeks) and adult Dutch Belted rabbits, but we have not attempted this method in younger animals or smaller breeds. In situations where the rabbit patient is small, or if a laryngoscope with a small Miller blade is unavailable, an alternative option is to use a handheld otoscope rather than the laryngoscope.15,20 Using an otoscope with a #5 ear speculum, the larynx is visualized, the guide catheter is placed by direct visualization, and the speculum is removed prior to proceeding following the same steps as outlined above.
It is important to use great care when the guide catheter is placed to avoid the possibility of causing trauma or bronchospasm as the end of the catheter passes into the trachea and touches the tracheal bifurcation. The risk of these complications can be reduced or avoided by feeding the catheter with very gentle movements, stopping as soon as there is resistance, and avoiding movement of the head and neck once the catheter is in place. In addition, when advancing the endotracheal tube over the catheter, there is the potential for the catheter to catch on the endotracheal tube connector. Thus, it is important to make sure that the catheter slides through the connector without catching; alternatively, the connector can be removed and replaced when securing the endotracheal tube or attaching to the anesthetic circuit.
With any method of intubation, there is the potential for complications and adverse effects. Using the direct visualization method with a laryngoscope, improper insertion of the laryngoscope blade or aggressive placement of the endotracheal tube can result in laryngeal, tongue, or tooth injuries. Correct positioning of the animal is the first critical step in minimizing trauma during visualization of the larynx. The spine should be straight, with the head in direct alignment with the spine; if the animal is not straight, or if the neck is twisted to the left or right, this will make visualization of the larynx more difficult and may result in excessive trauma with the laryngoscope blade. Another critical key to success when using this technique is to follow the roof of the mouth towards the larynx, and once the soft palate is visualized, shift the blade into a proper position by bending at the wrist. This keeps the head in place, while moving the tip of the blade into optimal viewing position of the vocal cords, which are located quite ventral in the rabbit20,24.
Because direct visualization of the larynx can be tricky if the precise techniques we outline above are not followed, the use of alternative intubation strategies that do not require visualization of the larynx can be appealing. For example, the blind technique is one possible option that does not require use of a laryngoscope16,17,18,19,20. However, use of this technique relies on the ability to detect airflow through the endotracheal tube in order to direct the placement of the tube, and if the animal&#39;s breaths are very shallow or arrested, use of this method is not reliable. Supraglottic devices may also be considered, since placement of these devices does not require visualization of the larynx and they are generally very easy to place. These devices do not directly enter the trachea, however, and thus there is potential for lack of direct flow to the airway, inability to ventilate if laryngospasm develops, lack of a seal tight enough for ventilation during open-chest procedures or emergency resuscitation, and development of gastric tympany3,4,8. Newer devices specifically designed for rabbits may offer a promising alternative, but more studies are needed to evaluate these devices, and the concerns regarding lack of entry into the trachea still remain. Gastric tympany can also be a potential complication in an intubated animal if the endotracheal tube is accidentally placed into the esophagus; however, use of a direct visualization technique minimizes the risk of this complication by verifying proper placement of the guide catheter into the larynx.
Trauma to the arytenoid and/or tracheal mucosa is another potential concern when inserting an endotracheal tube, particularly when using inflatable cuffed tubes26,27,28. The trachea in rabbits may be easily injured, which can lead to complications such as subcutaneous emphysema, pneumothorax, and even death10,27,28,29. While performing the technique we describe here, care should be taken to prevent tracheal injury by use of uncuffed endotracheal tubes, ensuring the rounded tip of the guide catheter is inserted into the trachea, and not forcing the guide catheter into the glottis. In addition, selection of an appropriately sized endotracheal tube is of critical importance. Endotracheal tube size should be considered against the size of the animal, as well as compared to the opening of the laryngeal inlet since it is smaller in diameter than the trachea. An endotracheal tube too large in diameter may result in trauma to the arytenoid mucosa or cartilage. On the other hand, use of a smaller endotracheal tube results in increased airway resistance and increases the chances of obstruction due to mucus or a kink. For this reason, respiratory monitoring is very important in intubated rabbits. In addition to pulse oximetry, measurement of airflow should be monitored with capnography; if a capnograph is unavailable, airway monitors are an alternative that can be used to detect flow of air (but do not provide measurements of exhaled carbon dioxide). In addition, provision of assisted ventilation can be valuable, particularly in cases when smaller diameter tubes are used24,30.
In addition to the already mentioned benefits of using this technique, another major advantage of this method is that it can be performed by a single individual without the need for an assistant, which can be important in a busy clinical setting. In this situation, it is important to gather all necessary supplies and have them within easy reach during the intubation process, particularly when the guide catheter is in place and movement of the head should be avoided.
The technique we describe here is used by the veterinary and research staff in our institution for all rabbit intubations, which amounts to approximately 200-400 intubations per year. It is very easily taught in a typical rabbit anesthesia training session, which takes approximately 30 minutes. We have found that it is relatively easy to learn, with students learning it within their first 3 attempts and generally able to repeat the process successfully in the next additional attempt.For future study, it would be valuable to compare time to proficiency and ability to maintain proficiency over time for this method compared to other intubation techniques. While this has been studied for variations of blind techniques16, a comprehensive assessment of the learning curve for a wide variety of intubation techniques has not yet been performed. Another potential future direction of study could include comparing direct endotracheal intubation to the newer species-specific supraglottic airway devices, particularly concerning function during positive pressure ventilation.
In conclusion, the technique for intubation of rabbits we describe here is non-invasive, reliable, does not require the use of specialized equipment, is easy to master, and can be performed by a single individual without the need for an assistant. All airway access techniques present some level of risk, but by using a direct visualization technique and following the steps we describe above to minimize complications, the risks are far outweighed by the benefits of having an adequate airway during anesthesia.

Rabbits have a narrow, deep oral cavity that restricts visibility of the larynx for endotracheal intubation. They also have a very large fleshy tongue and cheek teeth which, when combined with the narrow oral cavity, can cause issues when inserting a laryngoscope and visualizing the glottis to insert an endotracheal tube1. Great care must be taken to avoid causing injury to the oral cavity and airway when attempting intubation. The laryngeal tissues are very friable and excessive trauma from repeated attempts to intubate can cause significant inflammation, making intubation more difficult2.
For these reasons, a variety of alternative techniques have been described, most notably supraglottic airway devices including the laryngeal tube3,4, the laryngeal mask airway (LMA)4,5,6 and a species-specific supraglottic airway device7. A laryngeal tube consists of a tube with two balloons that direct air flow into the trachea: a larger balloon in the middle of the tube forms a seal in the upper pharynx and a smaller balloon at the distal end seals the esophageal inlet3. The LMA consists of a tube with an elliptical mask at the distal end that is inflated to form an airtight seal over the glottis4,5,6. The species-specific device is the first veterinary species-specific supraglottic airway device, currently available for cats and rabbits, designed to fit the anatomical structures for each species to create a pressure seal in the pharynx7. However, because these devices do not directly enter the trachea, they may present risks for complications such as gastric tympany. The lack of a proper seal within the trachea presents a concern regarding the adequacy for use of these devices for positive pressure ventilation for respiratory support, emergencies, or thoracic surgery3,8. In addition, lack of direct intubation of the trachea may result in personnel exposure to waste anesthetic gases5.
Many methods for direct endotracheal intubation have been described9,10,11,12,13,14,15. One popular method is the blind technique, which may decrease the chance of injury to the oropharyngeal cavity since insertion of a laryngoscope isn&#39;t necessary16,17,18,19,20. As a visually blind technique, this method requires monitoring for air flow through the endotracheal tube and careful manipulation to guide placement. Although this method is commonly used in research and clinical veterinary practice, in the authors&#39; experience, this technique can be time consuming and not always successful, and thus is more difficult to teach and may require extra dosing of anesthetic agents9,10. In addition, because monitoring for respiration is used to guide proper placement of the tube, this method would be more difficult and less reliable during respiratory arrest. Another popular method is to use a traditional laryngoscope technique to intubate as is done with larger species. However, the traditional laryngoscope technique in rabbits is difficult in that the view of the vocal cords is obstructed when the endotracheal tube is inserted into the mouth. Therefore, use of this method can result in repeated intubation attempts, increased anesthetic dosing, and injury to the oral cavity and airway10.
More expensive and technically challenging options for intubation also exist. The fiberoptic laryngoscope allows easy insertion of the endotracheal tube by inserting the laryngoscope into the tube and visualizing the insertion into the trachea on a screen. Similarly, endoscopes can be used to visualize the laryngeal area and guide placement of the tube. However, both of these methods require the use of expensive equipment that may not be readily available in typical clinical settings9,11,12. Intubation via the nasal cavity is an alternative method that has been described, though this presents a risk of introducing opportunistic pathogens from the nasal cavity into the lungs, particularly in rabbits that are not specific-pathogen-free21. Lastly, more invasive methods of intubation have also been published, including retrograde insertion of a guide catheter through a needle placed percutaneously into the trachea, as well as the use of a needle cricothyroidotomy followed by placement of a guide wire, vessel dilator, and sheath introducer13,14. As invasive techniques, these methods have potential complications such as post-procedural pain and infection. The technique we describe here, using a laryngoscope and guide catheter, can be taught with minimal training with a short time to proficiency, is non-invasive, can be performed by a single individual without the need for an assistant, and does not require the use of specialized equipment.

<table>
	<tbody>
		<tr>
			<td>Ketalar 
			Ketamine HCl Inj.</td>
			<td>JHP Pharmaceuticals</td>
			<td>NDC 42023-113-10</td>
			<td>35mg/kg IM in epaxial muscles or quadriceps muscle (preferred)</td>
		</tr>
		<tr>
			<td>Anased 
			Xylazine (100mg/ml)</td>
			<td>Lloyd Labs</td>
			<td>NADA #139-236</td>
			<td>5mg/kg IM in epaxial muscles or quadriceps muscle (preferred)</td>
		</tr>
		<tr>
			<td>Welch Allyn Laryngoscope Handle</td>
			<td>Welch Allyn</td>
			<td>60300</td>
			<td>Requires 2 size C alkaline batteries</td>
		</tr>
		<tr>
			<td>Size 0 or 1 Miller Laryngoscope Blade</td>
			<td>Welch Allyn</td>
			<td>68040</td>
			<td></td>
		</tr>
		<tr>
			<td>2.5 &#8211; 4.0 mm ID uncuffed endotracheal tube</td>
			<td>Jorgenson</td>
			<td>J0149V</td>
			<td>Suitable for 2.5-5 kg rabbits 
			3.5 mm most commonly used</td>
		</tr>
		<tr>
			<td rowspan="2">Sovereign polypropylene catheter, 5 Fr x 22&#8221;</td>
			<td rowspan="2">Medtronic</td>
			<td rowspan="2">8890703310</td>
			<td rowspan="2">Cut to 8-10 inches 
			Mark the cut end with permanent marker to indicate which end not to insert</td>
		</tr>
		<tr>
			<td>Lidocaine 2% HCL Inj</td>
			<td>VetOne</td>
			<td>L-2000-04</td>
			<td></td>
		</tr>
		<tr>
			<td>4-Ply Nonsterile, Bulk Packed Versalon Sponges (Gauze pads)</td>
			<td>Covidien</td>
			<td>KC9024A</td>
			<td></td>
		</tr>
		<tr>
			<td>Tape, Twill, Bleached &#189;&#8221; x 72 yds</td>
			<td>Encompass Textiles</td>
			<td>48210-025</td>
			<td>Used for securing endotracheal tube</td>
		</tr>
		<tr>
			<td>HR Lubricating Jelly</td>
			<td>HR Pharmaceuticals</td>
			<td>201</td>
			<td>For lubricating endotracheal tube tip</td>
		</tr>
		<tr>
			<td>Great Plains Ballistics: BAAM (Beck Airway Airflow Monitor) 22</td>
			<td>BoundTree Medical</td>
			<td>21410</td>
			<td>For confirmation of successful intubation</td>
		</tr>
		<tr>
			<td>Ambu Pediatric Co2 Detector</td>
			<td>AMBU INC</td>
			<td>172713</td>
			<td>For confirmation of successful intubation</td>
		</tr>
	</tbody>
</table>

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.

At the authors&#39; institution, this technique is typically taught to laboratory and veterinary personnel during training sessions on rabbit handling and anesthesia techniques. Teaching sessions are performed with dedicated training animals. The trainer demonstrates the technique once and then trainees perform the technique under the guidance of the trainer. Trainees typically achieve success within 1-3 attempts. After they have successfully intubated the rabbit once, individuals are required to perform the intubation a second time, and are typically able to successfully intubate the rabbit again on the first additional attempt. Feedback during these training sessions, particularly from individuals who have been previously trained with alternative techniques, consistently includes comments on how easy this technique is to learn and complete successfully.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56369/56369fig1.jpg" /> 
Figure 1. Supplies for endotracheal intubation: 1) Laryngoscope with size 0 or 1 Miller laryngoscope blade, 2) umbilical tape (or other supplies for securing endotracheal tube), 3) 0.25 mL of 2% lidocaine, 4) 5-French polypropylene catheter cut to 8-10 inches with the non-rounded end marked with a permanent marker to clearly indicate which end should not be inserted into the trachea (arrow), 5) air flow detection device, 6) sterile lubricant to be applied to the distal end of the tube, 7) uncuffed endotracheal tube.&#160;<a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig1large.jpg" style="font-size: 14px;" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56369/56369fig2.jpg" /> 
Figure 2: Sternal positioning for endotracheal intubation. The rabbit is positioned sternally with the forelimbs extended to the end of the table. The hind limbs are extended back, and the rabbit&#39;s head is aligned with the spine. The left hand is used to elevate the head into extension in preparation for insertion of the laryngoscope. <a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56369/56369fig3.jpg" /> 
Figure 3: Insertion of laryngoscope. A) The laryngoscope is inserted into the left diastema using the right hand, following the roof of the mouth with the tip of the blade until the soft palate is visible. Once the soft palate is visible, the laryngoscope is tipped forward while extending the rabbit&#39;s head back and neck forward to visualize the larynx. B) Visualization of the larynx. <a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56369/56369fig4.jpg" /> 
Figure 4: Advancement of the guide catheter. A) After switching the laryngoscope blade to the left hand, the right hand advances the rounded tip of the guide catheter through the opening past the larynx. B) The guide catheter is passed through the vocal chords with direct visualization. <a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56369/56369fig5.jpg" /> 
Figure 5: Advancement of the tracheal tube. The endotracheal tube is threaded over the guide catheter and is advanced until there is resistance from the vocal cords. <a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56369/56369fig6.jpg" /> 
Figure 6: lidocaine instillation. Lidocaine (2%, 0.25 mL) is instilled into the opening of the endotracheal tube. <a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56369/56369fig7.jpg" /> 
Figure 7: Endotracheal tube advancement. Two minutes after instillation of the lidocaine, the endotracheal tube is advanced by gently twisting it to pass through the vocal cords. <a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/56369/56369fig8.jpg" /> 
Figure 8: Verification of air flow through the endotracheal tube. An airflow detection device may be used to verify air flow through the endotracheal tube, or a dental mirror may also be used to detect the condensation from breaths indicating successful intubation.&#160;<a href="https://cloudflare.jove.com/files/ftp_upload/56369/56369fig8large.jpg" style="font-size: 14px;" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Endotracheal Intubation of Rabbits Using a Polypropylene Guide Catheter at JoVE.com

Endotracheal Intubation of Rabbits Using a Polypropylene Guide Catheter

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 ...

endotracheal-intubation-rabbits-using-polypropylene-guide

Nanyang Technological University

Research

JoVE Journal

Biology

33.3K Views.  Mayo Clinic. 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.

Video: Endotracheal Intubation of Rabbits Using a Polypropylene Guide Catheter

Generation of Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing and adult tissue. An advance in this concept is Genetic Inducible Fate Mapping (GIFM), linking gene expression, cell fate, and cell behaviors in vivo, to create fate maps based on genetic lineage.
GIFM exploits X-CreER lines where X is a gene or set of gene regulatory elements that confers spatial expression of a modified bacteriophage protein, Cre recombinase (CreERT). CreERT contains a modified estrogen receptor ligand binding domain which renders CreERT sequestered in the cytoplasm in the absence of the drug tamoxifen. The binding of tamoxifen releases CreERT, which translocates to the nucleus and mediates recombination between DNA sequences flanked by loxP sites. In GIFM, recombination typically occurs between a loxP flanked Stop cassette preceding a reporter gene such as GFP.
Mice are bred to contain either a region- or cell type-specific CreER and a conditional reporter allele. Untreated mice will not have marking because the Stop cassette in the reporter prevents further transcription of the reporter gene. We administer tamoxifen by oral gavage to timed-pregnant females, which provides temporal control of CreERT release and subsequent translocation to the nucleus removing the Stop cassette from the reporter. Following recombination, the reporter allele is constitutively and heritably expressed. This series of events marks cells such that their genetic history is indelibly recorded. The recombined reporter thus serves as a high fidelity genetic lineage tracer that, once on, is uncoupled from the gene expression initially used to drive CreERT.
We apply GIFM in mouse to study normal development and ascertain the contribution of genetic lineages to adult cell types and tissues. We also use GIFM to follow cells on mutant genetic backgrounds to better understand complex phenotypes that mimic salient features of human genetic disorders.
This video article guides researchers through experimental methods to successfully apply GIFM. We demonstrate the method using our well characterized Wnt1-CreERT;mGFP mice by administering tamoxifen at embryonic day (E)8.5 via oral gavage followed by dissection at E12.5 and analysis by epifluorescence stereomicroscopy. We also demonstrate how to micro-dissect fate mapped domains for explant preparation or FACS analysis and dissect adult fate-mapped brains for whole mount fluorescent imaging. Collectively, these procedures allow researchers to address critical questions in developmental biology and disease models.

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Genetic Inducible Fate Mapping (GIFM) marks and tracks cells with fine spatial and temporal control in vivo and elucidates how cells from a specific genetic lineage contribute to developing and adult tissues. Demonstrated here are the techniques required to fate map E12.5 mouse embryos for epifluorescent and explant analysis.

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing ...

Rapid PCR Thermocycling using Microscale Thermal Convection

Many molecular biology assays depend in some way on the polymerase chain reaction (PCR) to amplify an initially dilute target DNA sample to a detectable concentration level. But the design of conventional PCR thermocycling hardware, predominantly based on massive metal heating blocks whose temperature is regulated by thermoelectric heaters, severely limits the achievable reaction speed1. Considerable electrical power is also required to repeatedly heat and cool the reagent mixture, limiting the ability to deploy these instruments in a portable format.
Thermal convection has emerged as a promising alternative thermocycling approach that has the potential to overcome these limitations2-9. Convective flows are an everyday occurrence in a diverse array of settings ranging from the Earth's atmosphere, oceans, and interior, to decorative and colorful lava lamps. Fluid motion is initiated in the same way in each case: a buoyancy driven instability arises when a confined volume of fluid is subjected to a spatial temperature gradient. These same phenomena offer an attractive way to perform PCR thermocycling. By applying a static temperature gradient across an appropriately designed reactor geometry, a continuous circulatory flow can be established that will repeatedly transport PCR reagents through temperature zones associated with the denaturing, annealing, and extension stages of the reaction (Figure 1). Thermocycling can therefore be actuated in a pseudo-isothermal manner by simply holding two opposing surfaces at fixed temperatures, completely eliminating the need to repeatedly heat and cool the instrument.
One of the main challenges facing design of convective thermocyclers is the need to precisely control the spatial velocity and temperature distributions within the reactor to ensure that the reagents sequentially occupy the correct temperature zones for a sufficient period of time10,11. Here we describe results of our efforts to probe the full 3-D velocity and temperature distributions in microscale convective thermocyclers12. Unexpectedly, we have discovered a subset of complex flow trajectories that are highly favorable for PCR due to a synergistic combination of (1) continuous exchange among flow paths that provides an enhanced opportunity for reagents to sample the full range of optimal temperature profiles, and (2) increased time spent within the extension temperature zone the rate limiting step of PCR. Extremely rapid DNA amplification times (under 10 min) are achievable in reactors designed to generate these flows.

We describe a novel method to perform DNA replication via the polymerase chain reaction (PCR). Thermal convection is harnessed to continuously shuttle reagents between denaturing, annealing, and extension conditions by maintaining opposing surfaces of the reactor at constant temperature. This inherently simple design promises to make rapid PCR more accessible.

Many molecular biology assays depend in some way on the polymerase chain reaction (PCR) to amplify an initially dilute target DNA sample to a detectable ...

A Practical Guide to Phylogenetics for Nonexperts

Many researchers, across incredibly diverse foci, are applying phylogenetics to their research question(s). However, many researchers are new to this topic and so it presents inherent problems. Here we compile a practical introduction to phylogenetics for nonexperts.&#160;We outline in a step-by-step manner, a pipeline for generating reliable phylogenies from gene sequence datasets. We begin with a user-guide for similarity search tools via online interfaces as well as local executables. Next, we explore programs for generating multiple sequence alignments followed by protocols for using software to determine best-fit models of evolution. We then outline protocols for reconstructing phylogenetic relationships via maximum likelihood and Bayesian criteria&#160;and finally describe tools for visualizing phylogenetic trees. While this is not by any means an exhaustive description of phylogenetic approaches, it does provide the reader with practical starting information on key software applications commonly utilized by phylogeneticists. The vision for this article would be that it could serve as a practical training tool for researchers embarking on phylogenetic studies&#160;and also serve as an educational resource that could be incorporated into a classroom or teaching-lab.

Here we describe a step-by-step pipeline for generating reliable phylogenies from nucleotide or amino acid sequence datasets. This guide aims to serve researchers or students new to phylogenetic analysis.&#160;

Many researchers, across incredibly diverse foci, are applying phylogenetics to their research question(s). However, many researchers are new to this ...

Determination of Protein-ligand Interactions Using Differential Scanning Fluorimetry

A wide range of methods are currently available for determining the dissociation constant between a protein and interacting small molecules. However, most of these require access to specialist equipment, and often require a degree of expertise to effectively establish reliable experiments and analyze data. Differential scanning fluorimetry (DSF) is being increasingly used as a robust method for initial screening of proteins for interacting small molecules, either for identifying physiological partners or for hit discovery. This technique has the advantage that it requires only a PCR machine suitable for quantitative PCR, and so suitable instrumentation is available in most institutions; an excellent range of protocols are already available; and there are strong precedents in the literature for multiple uses of the method. Past work has proposed several means of calculating dissociation constants from DSF data, but these are mathematically demanding. Here, we demonstrate a method for estimating dissociation constants from a moderate amount of DSF experimental data. These data can typically be collected and analyzed within a single day. We demonstrate how different models can be used to fit data collected from simple binding events, and where cooperative binding or independent binding sites are present. Finally, we present an example of data analysis in a case where standard models do not apply. These methods are illustrated with data collected on commercially available control proteins, and two proteins from our research program. Overall, our method provides a straightforward way for researchers to rapidly gain further insight into protein-ligand interactions using DSF.

Differential scanning fluorimetry is a widely used method for screening libraries of small molecules for interactions with proteins. Here, we present a straightforward method to extend these analyses to provide an estimate of the dissociation constant between a small molecule and its protein partner.

A wide range of methods are currently available for determining the dissociation constant between a protein and interacting small molecules. However, most ...

A Guide to Modern Quantitative Fluorescent Western Blotting with Troubleshooting Strategies

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins within complex biological samples. Since then, western blotting methodology has become a common component of the molecular biologists experimental repertoire. A cursory search of PubMed using the term &#8220;western blot&#8221; suggests that in excess of two hundred and twenty thousand published manuscripts have made use of this technique by the year 2014. Importantly, the last ten years have seen technical imaging advances coupled with the development of sensitive fluorescent labels which have improved sensitivity and yielded even greater ranges of linear detection. The result is a now truly Quantifiable Fluorescence based Western Blot (QFWB) that allows biologists to carry out comparative expression analysis with greater sensitivity and accuracy than ever before. Many &#8220;optimized&#8221; western blotting methodologies exist and are utilized in different laboratories. These often prove difficult to implement due to the requirement of subtle but undocumented procedural amendments. This protocol provides a comprehensive description of an established and robust QFWB method, complete with troubleshooting strategies.

The advancement of western blotting using fluorescence has allowed detection of subtle changes in protein expression enabling quantitative analyses. Here we describe a robust methodology for detection of a range of proteins across a variety of species and tissue types. A strategy to overcome common technical problems is also provided.

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins ...

Discrimination and Characterization of Heterocellular Populations Using Quantitative Imaging Techniques

Cellular processes are complex and result from the interplay between multiple cell types and their environment. Existing cell biology techniques often do not allow for accurate interpretation of this interplay. Using a quantitative imaging-based approach, we present a high-content protocol for characterizing the dynamic phenotypic responses (i.e. morphology changes, proliferation, apoptosis) of heterogeneous cell populations to changes in environmental stimuli. We highlight our ability to distinguish between cell types based upon either fluorescence intensity or inherent morphology features depending on the application. This platform allows for a more comprehensive characterization of subpopulation response to perturbation while utilizing shorter time, smaller amounts of reagents, and lower likelihood of error than traditional cell biology assays. However, in some cases, cell populations may be difficult to identify and quantitate based on complex cellular features and will require additional troubleshooting; we highlight some of these circumstances in the protocol. We demonstrate this application using response to drug in a cancer model; however, it can easily be applied more broadly to other physiological processes. This protocol allows one to identify subpopulations within a co-culture system and characterize the particular response of each to external stimuli.

We have developed a novel protocol for studying heterocellular population dynamics in response to perturbations. This manuscript describes an imaging-based platform that produces quantitative datasets for simultaneous characterization of multiple cellular phenotypes of heterocellular populations in a robust manner.

Cellular processes are complex and result from the interplay between multiple cell types and their environment. Existing cell biology techniques often do ...

A Versatile Model of Hard Tick Infestation on Laboratory Rabbits

The use of live animals in tick research is crucial for a variety of experimental purposes including the maintenance of hard tick colonies in the laboratory. In ticks, all developmental stages (except egg) are hematophagous, and acquiring a blood-meal when attached to their vertebrate hosts is essential for the successful completion of their life cycle. Here we demonstrate a simple method that uses easily openable capsules for feeding of hard ticks on rabbits. The advantages of the proposed method include its simplicity, short duration and most importantly versatile adjustment to the needs of specific experimental requirements. The method makes possible the use of multiple chambers (of various sizes) on the same animal, which permits feeding of multiple stages or different experimental groups while reducing the overall animal requirement. The non-irritating and easily accessible materials used minimizes discomfort to the animals, which can be easily recovered from an experiment and offered for adoption or reused if the ethical protocol allows it.

We have developed a simple and versatile system to feed hard ticks on laboratory rabbits. Our non-laborious protocol uses easily accessible materials and can be adjusted depending on the requirements of the various experimental settings. The method allows comfortable monitoring and/or sampling of ticks during the entire feeding period.

The use of live animals in tick research is crucial for a variety of experimental purposes including the maintenance of hard tick colonies in the ...

A Sectioning, Coring, and Image Processing Guide for High-Throughput Cortical Bone Sample Procurement and Analysis for Synchrotron Micro-CT

Bone is a dynamic and mechanically active tissue that changes in structure over the human lifespan. The products of the bone remodeling process have been studied substantially using traditional two-dimensional techniques. Recent advancements in X-ray imaging technology via desktop micro-computed tomography (&#181;CT) and synchrotron radiation micro-computed tomography (SR&#181;CT) have allowed for the acquisition of high-resolution three-dimensional (3D) scans of a larger field of view (FOV) than other 3D imaging techniques (e.g., SEM) providing a more complete picture of microscopic structures within human cortical bone. The specimen should be accurately centered within the FOV, however, to limit the appearance of streak artifacts known to impact data analysis. Previous studies have reported procurement of irregularly shaped rectilinear bone blocks that result in imaging artifacts due to uneven edges or image truncation. We have applied a geological sampling protocol (coring) to procure consistently sized cortical bone core specimens for SR&#181;CT experiments from the anterior aspect of human femora. This coring method is efficient and minimally destructive to tissue. It creates uniform cylindrical samples that decrease imaging artifacts by nature of being isometric during rotation and providing a uniform path length for X-ray beams throughout scanning. Image processing of X-ray tomographic data of cored and irregularly shaped samples confirms the potential of the technique to improve visualization and analysis of cortical bone microarchitecture. A goal of this protocol is to deliver a reliable and repeatable method for the extraction of cortical bone cores that is adaptable for various types of high-resolution bone imaging experiments. An overarching goal of the work is to create a standardized cortical bone procurement for SR&#181;CT that is affordable, consistent, and straightforward. This procedure may further be adapted by researchers in related fields who commonly evaluate hard composite materials such as in biological anthropology, geosciences, or material sciences.

We employed a geological (coring) sampling protocol to procure cortical bone specimens of uniform size for SR&#181;CT experiments from the anterior aspect of human femora. This method is minimally destructive, efficient, results in cylindrical specimens that minimize imaging artifacts from irregular sample shapes and improves microarchitectural visualization and analysis.

Bone is a dynamic and mechanically active tissue that changes in structure over the human lifespan. The products of the bone remodeling process have been ...

A Step-by-Step Guide to Mosquito Electroantennography

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when acquiring a blood-meal. To locate a host to feed on, mosquitoes rely on a wide range of sensory cues, including visual, mechanical, thermal, and olfactory. The study details a technique, electroantennography (EAG), that allows researchers to assess whether the mosquitoes can detect individual chemicals and blends of chemicals in a concentration-dependent manner. When coupled with gas-chromatography (GC-EAG), this technique allows to expose the antennae to a full headspace/complex mixture and determines which chemicals present in the sample of interest, the mosquito can detect. This is applicable to host body odors as well as plant floral bouquets or other ecologically relevant odors (e.g., oviposition sites odorants). Here, we described a protocol that permits long durations of preparation responsiveness time and is applicable to both female and male mosquitoes from multiple genera, including Aedes, Culex, Anopheles, and Toxorhynchites mosquitoes. As olfaction plays a major part in mosquito-host interactions and mosquito biology in general, EAGs and GC-EAG can reveal compounds of interest for the development of new disease vector control strategies (e.g., baits). Complemented with behavioral assays, the valence (e.g., attractant, repellent) of each chemical can be determined.

The present article details a step-by-step protocol for successful and low noise electroantennograms in several genera of mosquitoes, including both females and males.

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when ...