Improved Methodology for Liquid Delivery to the Mouse Lung: Intubation using a Consumer Otoscope

[Announcer] Improved methodology for liquid delivery to the mouse lung. Intubation using a consumer otoscope. All animal procedures and this content were reviewed by the Tulane University Department of Comparative Medicine and are approved on Protocol 1810. Seek IACUC approval and consult veterinary staff at your institution before continuing. The development of new pulmonary therapeutics is dependent on reliable, repeatable delivery to the mouse lung but has traditionally relied on intranasal administration, oropharyngeal aspiration by tongue full or surgical cutdown. The following method has reduced variability compared to tongue full, allowing for smaller animal cohorts. It is also far less invasive than cutdown. Procedure, step one, setup. The rodent intubation kit initially used ships with intubation safety wedges. These are small conical sleeves that fit over a 20-gauge, one-inch flexible catheter. They prevent the catheter from damaging the carina. Clean a safety wedge with an isopropanol prep pad before using it. Slide the clean safety wedge, green here, onto the 20 gauge flexible cannula. Gather the other tools needed including a commercial otoscope, mobile phone, mouse stand with nose cone, bent, finely serrated forceps, the endotracheal tube, and the fiber optic light, if using. Clean all of these with an isopropanol prep pad before use. Take an approximately 20 centimeter length of flexible tubing. Bend it in a U-shape and add a lure lock connection on one end. Secure the U-shape with a twist tie. Add two to 300 microliters of sterile filtered water or India ink to the tube. Hereafter, this assembly is referred to as a bubble tube. Take a suture loop by tying 13 centimeters of size zero rated silk suture in a circle. Place this loop at the top of the mouse stand and ensure both sides of the suture are well secured. Place the mouse stand on top of the metal magnifying light base. Position the helping hands around where the mouse's mouth will be. Place a mouse in the induction chamber. Use constant flow rate oxygen at 0.8 liters per minute and 2% isoflurane. Anesthetize the mouse sufficiently as to not awaken during placement of the nose cone. Do not significantly suppress the mouse's respiratory rate. Regular breaths aid in appreciation of the vocal folds. Connect the otoscope with the mobile phone and ensure the video feed is stable. Caution, to minimize the researcher's isofluorane exposure during this procedure, use a chemical respirator if conducting multiple intubations or training in this method. Alternatively, conduct the procedure in a few wood. Step two, animal staging. Take the first animal from the induction chamber as soon as they're asleep. Scruff them and place their top teeth on the length of suture. Apply light tension to the animal to maintain the positioning of the suture while laying the mouse back onto the stand. Adjust the neck angle slightly back, placing the neck in minor extension five to 15 degrees. Gently push the nose cone over the nares. Do not push the nose cone down firmly as it can interfere with the suture suspending the mouse. Using large, finely serrated bent forceps, firmly grasp the tongue as close to the bottom lip as possible. Firmly pull it down and away from the top teeth and over to the side of the bottom teeth. Caution, prolonged traction, excessive tension or crushing the tongue can compromise animal health. Please monitor for signs of trauma including bruising, bleeding, swelling, or desiccation of the tongue. If any are observed, abort the procedure and alert veterinary staff. If using helping hands, support the forceps in this position. Feed the fiber optic light if using through the flexible catheter with the intubation safety wedge in place. Step three, visual appreciation of the vocal cords and intubation. With the otoscope in your non-dominant hand, lightly press on the tongue. Then angle the camera and its light source to visualize the vocal cords near the base of the tongue. Due to slight variations in anatomy and tissue color, use the movement associated with breathing to confirm structural identification. When first beginning, orienting oneself in the video feed may prove difficult. These are the false vocal folds. These are the true vocal chords. This is reflection off of the buccal tissue. This is the probe and fiber optic light. This is the epiglottis. This is the soft palate. This is the rima glottidis, your target for intubation. This is the otoscope lip, depressing the tongue. This is the no-go region. With the fiber optic light on, if using, feed the cannula alongside the otoscope, making light contact with the base of the tongue or the lip portion of the otoscope. For reference, adjust the angle of approach for the catheter to be very shallow. Immediately after inserting the catheter, less than one millimeter beyond the vocal folds, raise the tip of the catheter so it is nearly parallel to the bench top. This prevents the epiglottis from forcing the catheter dorsally into the esophagus. Note, there should be almost no resistance when advancing the catheter. If resistance is encountered, immediately remove the catheter and allow respirations to return to normal. If bleeding is observed, consult veterinary staff. Step four, confirmation of endotracheal tube placement. In addition to visual confirmation that the tube is in the correct location using the otoscope, the trachea provides less resistance during the placement than the esophagus. The optical insertion depth is about five millimeters beyond the vocal folds. The light can be seen through the neck as a secondary method of confirming that the depth of insertion is appropriate. Note the intubation safety wedge should prevent excessive depth in endotracheal tube placement. The safety wedge will prevent trauma to the corina, however it need not be preventing advancement. Then, while holding the endotracheal tube in place, remove the fiber optic light. Confirm appropriate placement of the catheter using the bubble tube method. Secure the bubble tube in place using the lower lock. The tidal volume should be immediately apparent. If not, the endotracheal tube is misplaced. Remove it quickly and allow the mouse's normal respiratory rate to return. Step five, intratracheal delivery. Much like the tongue pull method for oropharyngeal delivery, delivery via intubation risks drowning the animal. In adolescent and adult mice, 50 microliters is a generally safe volume of fluid to deliver and is sufficient to spread throughout the lungs. If using a pipette for administration, pipette the desired volume directly into the catheter and allow the mouse to inhale the solution. To ensure diffuse delivery to the lungs, plus the delivered liquid into lungs with up to 500 microliters of air. If desired, multiple depressions of the pipeter can be conducted without risking negative aspiration of the liquid delivered. Note, use of more than the tidal volume, approximately 200 microliters, may be necessary when flushing the inoculum with air. However, the use of 500 microliters does not risk trauma as it is only about half of the vital capacity. Step six, assessment of delivery. A handheld fiber optic light can be made from cheaply and easily obtained components. A commercial fiber optic assembly, as shown here, may be used. However, a 50 ml conical tube lid with a hole punched in it using a 20-gauge needle through which a .75 millimeter plastic fiber optic cable is fed suffices. Both commercial and handmade devices were used in this video. Representative results, intratracheal delivery using intubation results in improved delivery and reduced variability when compared to oropharyngeal aspiration by tongue pull. Direct comparison of tongue pull and guided intubation. Three different investigators with buried levels of experience or asked to independently deliver 50 microliters of dilute India ink oropharyngeally by tongue pull to 10 to 12 week old C57 BL/6J mice of both sexes. Two researchers also delivered the same stock and volume of ink to the lung via intubation as described. The lungs were removed shortly after delivery and photographed under 2x magnification. Photographs were assessed for area stained by an independent blinded researcher using Image J. She found that more lung area was stained if intubation was used. Further, the variability, expressed as percent deviation from the mean, was reduced if intratracheal delivery following intubation was used rather than tongue pull. This station used for guided intubation is prepared as shown. From left, pipettes for delivery, bubble tube for confirmation of appropriate endotracheal tube placement, forceps, spatula, pediatric nasal speculum, helping hands, magnifying light, mouse stand with nose cone, fiber optic probe and anesthesia induction chamber. Intubation yields greater delivery to the lung with less variability relative to OP aspiration. 10-week-old female valve C mice were dosed with one times 10 to the 11th VG of AIV 6.2 luciferase and assessed with in vivo bioluminescent imaging seven days later. 10 minutes prior to imaging, they were injected subcutaneously with Luciferin. Representative a-vis images of mice receiving AV 6.2 luciferase by OP aspiration following tongue pull or intratracheal installation by intubation are shown. Oropharyngeal aspiration by tongue pull led to distribution of the signal in various parts of the upper airway, whereas intubation led to concentrated signal in the lung. Regions of interest are the same area, and average radiance within the region is shown on the image. Quantification of luciferase signals revealed improved delivery efficiency to the lungs by intubation and further emphasized the reduced variability in delivery using this method. Conclusion, as seen in the previous figures, interpulmonary inoculation using the traditional tongue pull method shows patchy, inconsistent inoculum distribution and greater variability in how much reaches the lungs. In contrast, the intubation method led to more consistent delivery to both the right and left lungs and was less variable. The robustly demonstrated positive outcomes of this intubation method are, one, replicable installation of up to 50 microliters to the mouse lungs, two, diffuse distribution of instilled liquid throughout the lungs, and three, reduced spread of the inoculum beyond the lungs. This method is fast, safe, and can be incorporated into laboratory procedures without costly equipment or time-intensive training.