This protocol describes a serial transoral laryngoscopy approach for mice and rats that permits close-up, unobstructed video imaging of the larynx during breathing and swallowing using an optimized anesthetic regimen and finely tuned endoscopic manipulation techniques.
The larynx is an essential organ in mammals with three primary functions – breathing, swallowing, and vocalizing. A wide range of disorders are known to impair laryngeal function, which results in difficulty breathing (dyspnea), swallowing impairment (dysphagia), and/or voice impairment (dysphonia). Dysphagia, in particular, can lead to aspiration pneumonia and associated morbidity, recurrent hospitalization, and early mortality. Despite these serious consequences, existing treatments for laryngeal dysfunction are largely aimed at surgical and behavioral interventions that unfortunately do not typically restore normal laryngeal function, thus highlighting the urgent need for innovative solutions.
To bridge this gap, we have been developing an experimental endoscopic approach to investigate laryngeal dysfunction in murine (i.e., mouse and rat) models. However, endoscopy in rodents is quite challenging due to their small size relative to current endoscope technology, anatomical differences in the upper airway, and the necessity for anesthesia to optimally access the larynx. Here, we describe a novel transoral laryngoscopy approach that permits close-up, unobstructed video imaging of laryngeal motion in mice and rats. Critical steps in the protocol include precise anesthesia management (to prevent overdosing that abolishes swallowing and/or risks respiratory distress-related mortality) and micromanipulator control of the endoscope (for stable video recording of laryngeal motion by a single researcher for subsequent quantification).
Importantly, the protocol can be performed over time in the same animals to study the impact of various pathological conditions specifically on laryngeal function. A novel advantage of this protocol is the ability to visualize airway protection during swallowing, which is not possible in humans due to epiglottic inversion over the laryngeal inlet that obstructs the glottis from view. Rodents therefore provide a unique opportunity to specifically investigate the mechanisms of normal versus pathological laryngeal airway protection for the ultimate purpose of discovering treatments to effectively restore normal laryngeal function.
The larynx is a cartilaginous organ located at the intersection of the respiratory and digestive tracts in the throat, where it functions as a valving mechanism to precisely control the flow and direction of air (i.e., during breathing and vocalizing) versus food and liquid (i.e., during swallowing). A wide range of disorders are known to affect the larynx, including congenital (e.g., laryngomalacia, subglottic stenosis), neoplastic (e.g., laryngeal papillomatosis, squamous cell carcinoma), neurological (e.g., idiopathic laryngeal paralysis, stroke, Parkinson's disease, amyotrophic lateral sclerosis), and iatrogenic (e.g., inadvertent injury during head or neck surgery). Regardless of the etiology, laryngeal dysfunction typically results in a symptom triad of dyspnea (difficulty breathing), dysphonia (voice impairment), and dysphagia (swallowing impairment) that negatively impact a person's economic and social welfare1,2,3,4.
Moreover, dysphagia, particularly in medically fragile individuals, can lead to aspiration pneumonia (due to food or liquid escaping through an incompletely closed larynx into the lungs) and associated morbidity, recurrent hospitalization, and early mortality5,6. Despite these serious consequences, existing treatments for laryngeal dysfunction are largely aimed at surgical and behavioral interventions that do not typically restore normal laryngeal function1,2,7,8,9,10, thus highlighting the urgent need for innovative solutions. Toward this goal, we have been developing an experimental endoscopic approach to investigate laryngeal dysfunction in murine (i.e., mouse and rat) models.
In human medicine, the gold standard for the evaluation of laryngeal dysfunction is endoscopic visualization, referred to as laryngoscopy11,12. Typically, a flexible endoscope is passed through the nose to examine the larynx, particularly the vocal folds and adjacent supraglottic and subglottic laryngeal structures. A rigid endoscope may also be used to visualize the larynx via the oral cavity. Either approach permits gross examination of laryngeal anatomy and can be used to assess laryngeal mobility and function during respiration, phonation, and a variety of airway protective reflexes such as coughing and the laryngeal adductor reflex13,14,15,16. During swallowing, however, the larynx is completely obscured by the epiglottis as it inverts to cover the laryngeal entrance, protecting it from the path of the food/liquid bolus being swallowed. As a result, direct visualization of laryngeal motion during swallowing is not possible in humans and must therefore be indirectly inferred using other diagnostic approaches (e.g., fluoroscopy, electromyography, electroglottography).
This paper describes an innovative laryngoscopy protocol for mice and rats that permits close-up, unobstructed imaging of breathing and airway protection during swallowing under light anesthesia. The protocol is compatible with a variety of commercially available endoscopy systems in combination with a custom platform to immobilize the anesthetized rodent throughout the procedure. Importantly, numerous designs/configurations of endoscopy platforms are indeed possible, depending on each lab's available resources and research agenda. Our intent here is to provide guidance for researchers to consider in the context of their research. Moreover, we aim to demonstrate how this laryngoscopy protocol can lead to a wealth of objective data that may spark novel insights into our understanding of laryngeal dysfunction and regeneration.
The combined effect of all the steps outlined in this murine laryngoscopy protocol results in a minimally invasive examination of the adult murine larynx that can be repeated in the same animals to detect and characterize laryngeal dysfunction over time in response to iatrogenic injury, disease progression, and/or treatment intervention relative to airway protection. Of note, this protocol does not evaluate vocalization-related laryngeal function.
The murine laryngoscopy protocol follows an approved Institutional Animal Care and Use Committee (IACUC) protocol and National Institutes of Health (NIH) Guidelines. It was developed for use with over 100 adult C57BL/6J mice and over 50 adult Sprague Dawley rats, approximately equal sexes and 6 weeks-12 months old for both species. Additional protocol development is necessary for adaptation to younger/smaller rodents. Animals were group housed (up to four mice or two rats per cage, based on sex and litter). The standard vivarium conditions included static caging with strict regulation of ambient temperature (20-26 °C), humidity (30%-70%), and standard 12 h light cycle. All animals received fresh enrichment materials (e.g., hut/pipe, dental treats, nestlet) at weekly cage changes. Unlimited access to food and water was provided, except during a short (up to 4-6 h) food restriction prior to anesthesia as described below. Veterinary and research staff monitored the animals every day.
1. Animal anesthesia that does not abolish swallowing
2. Transoral passage of the endoscope to visualize the larynx
3. Close-up, unobstructed video recording of laryngeal motion during breathing and evoked swallowing
NOTE: Synchronous electrophysiological recording of breathing, swallowing, and swallow-breathing coordination is also an option.
4. Anesthesia recovery
5. Objective quantification of laryngeal motion during breathing versus swallowing
Successful use of this murine laryngoscopy protocol results in close-up visualization of the larynx during spontaneous breathing and evoked swallowing under healthy and disease conditions, as shown in Figure 6. Moreover, this protocol can be repeated multiple times in the same rodents to permit investigation of laryngeal function/dysfunction over time. As shown in Figure 7, we successfully repeated this laryngoscopy protocol 6x over a 4-month timespan to investigate the spontaneous recovery pattern in a rat surgical model of RLN injury (data not yet published). Attempts to use ISO anesthesia instead of KX resulted in near abolishment of swallowing (Figure 8) in rodents undergoing direct electrical stimulation of the right superior laryngeal nerve to evoke swallowing, as described in our previous experiments31,32. This occurred with ISO as low as 2%; reducing ISO below this level resulted in the return of spontaneous movement and was therefore avoided. This confounding effect of ISO highlights the importance of anesthesia selection for the successful use of this protocol.
When endoscopic image quality is good, representative video clips of breathing and swallowing can be analyzed using motion tracking software, as shown in Figure 9. Representative outcome measures automatically generated by our custom laryngeal tracking software are listed in Table 1. Note that several breathing- and swallowing-related outcome measures were markedly different between baseline and post-RLN transection in the same representative rat. Whereas glottal angles during breathing were similar between baseline and post-RLN transection, ratios of right/left laryngeal motion amplitude (i.e., mean motion range ratio or MMRR) and frequency (open-close cycle ratio or OCCR) during breathing were lower following transection. Similarly, the swallow duration was shorter following RLN transection.
If synchronous electrophysiological recordings (e.g., respiratory pneumogram and genioglossus EMG) are acquired, several additional objective outcome measures are quantifiable for correlation with laryngoscopy data. Examples of electrophysiology-based outcome measures of interest to our research are summarized in Figure 10. We are currently developing algorithms for automated quantification of these outcome measures.
Figure 1: Murine endoscopy platform. (A) Sideand (B) topviews of the custom murine endoscopy platform are shown, with essential components labeled. Note the tabletop beneath the heating pad is size-adjustable. Shown here are the tabletop and heating pad sizes used with rats, which are easily removed to expose a mouse-sized tabletop that accommodates a smaller heating pad (not shown). A custom adapter secures an endoscope to a micromanipulator that is attached to the platform base. This strategic design allows the entire platform to be moved as a unit during the endoscopy procedure, without risking injury to the animal from inadvertent/uncontrolled endoscope motion. The micromanipulator permits gross and micro adjustments of the endoscope tip in multiple directions, including x (left/right), y (forward/back), z (up/down), as well as rotation around y (pitch) and z (yaw). Please click here to view a larger version of this figure.
Figure 2: Otoscope and custom sheath for murine laryngoscopy. (A) Disassembled components of a commercial otoscope and custom stainless-steel sheath with adapter for murine laryngoscopy. (B) When assembled, the otoscope tip extends 1 mm beyond the metal sheath but is adjustable up to 5 mm as needed. This strategic design facilitates the advancement of the narrow otoscope tip into the rodent's laryngeal inlet while the slightly larger diameter (2.4 mm) metal sheath sufficiently holds the velum and epiglottis open for optimal visualization of the entire larynx during breathing and swallowing. Please click here to view a larger version of this figure.
Figure 3: Minimally invasive electrophysiological recording during endoscopy. A respiratory sensor is taped to the rodent's abdomen; an EMG electrode is inserted through the skin into the genioglossus muscle of the tongue; and a ground electrode is inserted subcutaneously at the hip. This approach permits the investigation of swallowing, breathing, and swallow-breathing coordination in synchrony with endoscopy. Note the skin is shaved and cleaned/disinfected at the electrode insertion sites. Yellow star = aluminum foil wrapped around the electrode lead connection sites to improve the signal-to-noise ratio in the electrophysiological recordings. Please click here to view a larger version of this figure.
Figure 4: Transoral endoscopy to visualize the larynx from a distance. (A) After gently retracting the tongue with a light finger grip, the endoscope is inserted between the tongue and central incisors at the red star location (i.e., the same side as the retracted tongue to maintain anatomical alignment with the endoscope shaft). (B) As the endoscope is advanced past the hard palate, (C) the epiglottis and velum come into view. (D) To visualize the glottis, the velum and epiglottis must be "decoupled" by applying pressure against the surface of the velum (at the location of the black outlined star in image C). Please click here to view a larger version of this figure.
Figure 5: Close-up endoscopic visualization of the larynx. (A) The endoscope tip is gently guided via micromanipulator control between the decoupled velum and epiglottis (at the location of the black-outlined star). As the endoscope advances, (B) the larynx comes into view and the glottal space (yellow star) is centered in the camera field of view via micromanipulator adjustments. (C) Continued micromanipulator advancement of the endoscope results in visualization of the entire ventral-dorsal and lateral dimensions of the larynx. Abbreviations: VC = ventral commissure of the larynx (i.e., the ventral junction point between the vocal folds); DC = dorsal commissure of the larynx (i.e., the dorsal junction point between the arytenoids); VFs = vocal folds; A = arytenoid. Please click here to view a larger version of this figure.
Figure 6: Visualization of the murine larynx during breathing and swallowing. Representative endoscopic images depicting laryngeal motion during breathing and swallowing in an adult Sprague Dawley rat (A–C) before and (D–F) after surgical transection of the right RLN. Note that the resting posture of the larynx appears unchanged (D) following RLN injury compared to (A) baseline. (B,E) During maximum inspiration, laryngeal asymmetry becomes obvious following RLN injury. Instead of both arytenoids abducting to enlarge the glottal space (yellow star), (B) as shown at baseline, (E) the ipsilateral (right) arytenoid (black asterisk) and vocal fold appear immobilized throughout the respiratory cycle following RLN injury. Right-sided asymmetry is also evident during swallowing. (C) At baseline, the arytenoids approximate at midline during swallowing, leaving a small ventral glottal gap between the vocal folds. (F) Following RLN injury, the ipsilateral arytenoid and VF move paradoxically (i.e., in the same direction as the unaffected side, red arrow) during swallowing, leaving a large glottal gap (yellow star) extending from the ventral to posterior laryngeal commissures. (F) This image provides direct evidence of impaired laryngeal airway protection in a rat model of iatrogenic RLN injury. (C,F) Note the larynx moves closer to the endoscope during swallowing, as indicated by the epiglottis and velum no longer being visible in the camera field of view. Black arrows indicate the direction of normal laryngeal motion whereas the red arrow indicates paradoxical motion; yellow star = glottal space. Abbreviations: VFs = vocal folds; A = arytenoid; RLN = recurrent laryngeal nerve. Please click here to view a larger version of this figure.
Figure 7: Using serial laryngoscopy to investigate laryngeal dysfunction during breathing and swallowing in a rat model of iatrogenic RLN injury. A Likert scale ranging from -2 to +2 was used to estimate laryngeal motion distance and direction in eight adult Sprague-Dawley rats over a 4 month period. After baseline laryngoscopy, the rats underwent a surgical procedure to transect the right RLN, followed by serial laryngoscopy at 1 week post-surgery, then again at 1 month intervals from 1 to 4 months post-surgery. All eight rats survived the procedure, thus demonstrating the effectiveness of our anesthesia regimen for serial laryngoscopy. (A) Videos were analyzed in real time and frame-by-frame/slow motion to quantify laryngeal motion during breathing, where 0 = no motion, 1 = some motion, and 2 = normal motion distance of the affected (right) side compared to the intact (left) side. (B) For swallowing, the glottal gap size was estimated as follows: 0 = no reduction in the glottal gap size (i.e., no laryngeal airway protection), 1 = some glottal gap reduction (i.e., incomplete airway protection), and 2 = complete adduction of the arytenoids, with only a small ventral glottal gap between the vocal folds (i.e., complete airway protection). Negative values for breathing and swallowing indicate laryngeal motion in the opposite direction than expected (i.e., paradoxical). Note that following RLN injury, both breathing and swallowing were negatively affected. Interestingly, laryngeal airway protection was complete (albeit paradoxical) at the 1 WPS timepoint but worsened thereafter, ranging from no protection to incomplete protection. Abbreviations: WPS = week post-surgery; MPS = months post-surgery; RLN = recurrent laryngeal nerve. Please click here to view a larger version of this figure.
Figure 8: Swallowing inhibited by ISO in rodents. (A) Image of a rodent undergoing laryngoscopy under ISO anesthesia, with labeled components of the custom ISO delivery system designed for this purpose. A major caveat of this innovative approach is the risk of personnel exposure to ISO. (B) Another downside to this approach is ISO suppression of swallowing. This side-by-side boxplot and scatterplot summarizes unpublished data comparing the effect of ISO versus KX anesthesia in mice (9 per group) undergoing direct electrical stimulation of the right superior laryngeal nerve to evoke swallowing. Shown here is the number of swallows evoked during a 5 min trial consisting of 20 s trains of 20 Hz stimulation followed by 10 s of rest. Compared to KX, mice anesthetized with ISO (as low as 2%) had significantly fewer swallows (p < 0.001, independent samples t-test), and swallowing was even abolished in 4/9 mice. Similar findings emerged from non-surgical experiments with both mice and rats (data not shown). Abbreviations: ISO = isoflurane; KX = ketamine-xylazine. Please click here to view a larger version of this figure.
Figure 9: Objective quantification of murine laryngeal motion using tracking software. The same images from Figure 6 showing breathing versus swallowing in a rat at baseline versus post RLN injury are shown here, with laryngeal motion tracking lines added by our custom software. Tracking lines were manually added to the first video frame along the medial border of the arytenoids for automated tracking of left (blue line) versus right (red line) laryngeal motion in the remaining video frames. Corresponding laryngeal motion graphs generated by our custom software from 2.5 s video clips show individual left/right motion versus derived global laryngeal motion, with labels corresponding to (A,D) laryngeal resting posture, (B,E) maximum glottal gap during inspiration, and (C,F) glottic closure during swallowing. Note the paradoxical motion of the right side (red arrows) post RLN injury, as well as the large glottal gap shown in the corresponding derived global motion graph. Representative outcome measures are included in Table 1. Abbreviation: RLN = recurrent laryngeal nerve. Please click here to view a larger version of this figure.
Figure 10: Electrophysiology-based outcome measures for correlation with laryngoscopy data. (A) Electrophysiology recordings during breathing and swallowing are shown for a healthy rat. The top window shows a respiratory trace (from a respiratory sensor taped to the rodent's abdomen), the middle window shows EMG activity in the genioglossus muscle, and the bottom window shows the filtered EMG activity. Note the rhythmic respiratory and EMG pattern during breathing, which is interrupted during swallowing events. Swallow events are readily detected via jagged motion in the respiratory trace (black arrows) that is immediately followed by brief apnea (red asterisk). (B) An expanded window of the dashed rectangular box in A shows how several outcome measures are quantified from the electrophysiological recordings. (A) Note that during inspiration (yellow panels), the respiratory trace (top window) is delayed ~150 ms (blue double arrow) compared to EMG bursting activity, which highlights temporal differences between the two electrophysiological methods. Representative electrophysiology-based outcome measures include 1) inspiratory phase duration (i); 2) inter-respiratory-interval (ii, calculated via the respiratory and filtered EMG channels); swallow area under the curve (iii); and swallow apnea (iv; calculated via the respiratory and filtered EMG channels). Abbreviation: EMG = electromyography. Please click here to view a larger version of this figure.
Outcome Measures | Baseline | Post-RLN Injury | |
Breathing | Minimum glottal angle (degrees) | 34.5 | 34.6 |
Maximum glottal angle (degrees) | 52.9 | 49.9 | |
Average glottal angle (degrees) | 43.7 | 42.2 | |
Mean motion range ratio (MMRR) | 1.26 | 0.29 | |
Open close cycle ratio (OCCR) | 1 | 0.11 | |
Swallowing | Laryngeal adduction (ms) | 200 | 233 |
Glottic closure duration (ms) | 67 | 0 | |
Laryngeal abduction (ms) | 233 | 67 | |
Total swallow duration (ms) | 500 | 300 |
Table 1: Representative outcome measures automatically generated by custom laryngeal tracking software. Abbreviation: RLN = recurrent laryngeal nerve.
Supplemental text about the laryngoscopy platform. Please click here to download this File.
We have successfully developed a replicable murine-specific laryngoscopy protocol that permits close-up visualization of laryngeal motion during breathing and swallowing. Importantly, the protocol can be performed over time in the same animals to study the impact of various pathological conditions specifically on laryngeal function. This protocol was developed over the past decade and has undergone substantial modification and troubleshooting along the way. Anesthesia optimization was the greatest challenge to overcome to prevent overdosing that abolishes swallowing and/or risks respiratory distress-related mortality. We initially used ISO, which resulted in the abolishment of swallowing, excess saliva production (that obstructs endoscopic visualization), and risk of personnel exposure, which are considered serious contraindications against using ISO for this procedure. We, therefore, focused on KX because it is a commonly used rodent anesthetic33,34,35.
We started our protocol development with mice14,22,29,30,36 while using a sialendoscope because of its smaller shaft diameter (1.1 mm) compared to other potentially suitable endoscopes for this purpose. Importantly, the sialendoscope has a working channel, which we initially used to deliver air pulses to evoke/study the laryngeal adductor reflex14. However, we found the laryngeal adductor reflex was often diminished/abolished in mice and rats, most likely due to general anesthesia and/or inactivation of laryngeal/pharyngeal sensory receptors secondary to mucosal drying from repeated air pulse delivery. Though the laryngeal adductor reflex could not be reliably evoked in our studies, swallowing surprisingly persisted and was readily evoked by mechanical stimulation at/near the laryngeal inlet. For this reason, we switched our focus to endoscopic analysis of mechanically evoked swallowing.
In the process, we abandoned the semi-rigid sialendoscope that was prone to breaking and had insufficient lighting and image resolution to reliably visualize and analyze laryngeal motion. In the exploration of numerous alternative endoscopes, we ultimately settled on a specific otoscope that was suitable for laryngoscopy with both mice and rats. Based on our experience, the most essential feature when selecting a suitable endoscope for murine laryngoscopy is a shaft diameter of less than 2 mm that can transmit sufficiently bright light for high-quality video capture. Larger diameter endoscopes cannot readily pass through the laryngeal inlet in mice and rats for close-up visualization of laryngeal motion. Otoscopes are particularly ideal for this purpose, given their excellent light transmission, rigid/durable design, and relatively low cost compared to other types of endoscopes (e.g., sialendoscope, flexible endoscope). In addition, while manual control of the endoscope is an option in stable hands, we consider micromanipulator control to be an essential feature of this laryngoscopy protocol. Importantly, micromanipulator control of the endoscope allows for stable video recording of laryngeal motion by a single researcher for subsequent quantification. To date, we have successfully used this otoscope-based protocol with adult mice and rats. We suspect smaller diameter endoscope options will be essential to perform laryngoscopy with younger/smaller rodents.
A novel advantage of our laryngoscopy protocol is the ability to visualize airway protection during swallowing in rodents, which is not possible in humans due to epiglottic inversion over the laryngeal inlet that obstructs the glottis from view. Rodents therefore provide a unique opportunity to specifically investigate the mechanisms of normal versus pathological laryngeal airway protection for the ultimate purpose of discovering treatments to effectively restore normal laryngeal function. This unique capability of this murine laryngoscopy protocol is a major advantage over videofluoroscopy (i.e., the other "gold standard" test for dysphagia), which has failed to detect aspiration in the numerous rodent models of dysphagia that we have developed/identified thus far30,36,37,38,39,40. This negative VFSS-based finding can be attributed to several anatomical differences in the upper airway of rodents that are apparent via our transoral endoscopy approach. First, the rodent larynx is positioned high in the nasopharynx where it is concealed by a tightly coupled epiglottis and velum that creates a cul-de-sac oral cavity. Additionally, the epiglottis at rest is entrapped beneath a mucosal sheath overlying the velum. This anatomical configuration results in rodents being obligate nasal breathers; thus, oral breathing in awake rodents is a sign of respiratory morbidity. During swallowing in healthy rodents, however, the epiglottis slides out from the mucosal sheath and inverts over the laryngeal inlet as the larynx elevates further into the nasopharynx, out of the path of the bolus. These dynamic upper airway events can be directly visualized/assessed via laryngoscopy in healthy rodents and models of laryngeal dysfunction.
Importantly, we have shown that despite not aspirating during VFSS testing, rodent models (e.g., iatrogenic RLN injury) indeed show evidence of impaired laryngeal airway protection (i.e., incomplete glottal closure) via laryngoscopy that is translational to human patients with dysphagia-related aspiration. Thus, this murine laryngoscopy protocol provides a useful translational platform to specifically investigate mechanisms of airway protection and targeted treatments, which currently remain elusive. Achieving this goal will require further development/optimization of our current method, which utilizes the endoscope tip to provide uncalibrated mechanical stimulation of the laryngeal/pharyngeal mucosa to evoke swallowing. More rigorous, precisely controlled methods for evoking swallowing are currently being explored in our lab, including direct electrical stimulation of the superior laryngeal nerve32,41 and chemical (e.g., citric acid42) stimulation of the laryngeal/pharyngeal mucosa. An additional limitation of this protocol is the supine positioning of the rodents, which does not mimic awake and natural feeding behavior. Initial protocol development included prone positioning, which resulted in restricted mandibular motion while also limiting visibility of the oral cavity, markedly impeding endoscope passage. It is possible to visualize the larynx from a distance with the endoscope tip in the hypopharynx; however, this approach typically requires manual retraction of the epiglottis, velum, and/or tongue for enhanced visualization of the larynx. We have fabricated a variety of custom manual retraction devices for this purpose (e.g., modified otoscope specula, modified pipette tips). However, portions of the larynx typically remain obscured from view, and the retraction devices can restrict laryngeal motion, which may be mistaken as dysfunction. Moreover, recent additional features of the endoscopy platform (e.g., Trendelenburg tilt, and a cutout between the ear bars to accommodate jaw motion) may facilitate testing rodents in the prone position. Ear bars and supplemental heat are necessary features of the laryngoscopy protocol. Ear bars prevent the head from moving during transoral manipulation of the endoscope. A homeothermic heating system maintains body temperature between 36 °C and 38 °C to promote stable anesthesia and prevent hypothermia throughout the procedure.
Now that methodology exists to reliably video record laryngeal motion during breathing and swallowing in rodents, high-throughput quantification is an essential next step. Therefore, our video analysis efforts are ongoing to determine which outcome measure generated by our custom software can best distinguish healthy from disease conditions as well as detect changes over time in response to natural disease progression or treatment interventions. The top candidates will be the focus of subsequent machine learning approaches to accelerate video imaging analysis. Importantly, cases of suboptimal image quality (e.g., insufficient lighting, anatomical structures outside the field of view, excess secretions obscuring laryngeal structures, etc.) are currently not amenable to laryngeal tracking; however, this barrier may be overcome in the future via machine learning tools. Until then, careful selection of video frame sequences that meet the criteria for laryngeal tracking analysis (as described in protocol section 5 remains paramount.
The authors have nothing to disclose.
This work was funded in part by two NIH grants: 1) a multi-PI (TL and NN) R01 grant (HL153612) from the National Heart, Lung, and Blood Institute (NHLBI), and 2) an R03 grant (TL, DC0110895) from the National Institute on Deafness and Other Communication Disorders (NIDCD). Our custom laryngeal motion tracking software development was partially funded by a Coulter Foundation grant (TL & Filiz Bunyak). We thank Kate Osman, Chloe Baker, Kennedy Hoelscher, and Zola Stephenson for providing excellent care of our laboratory rodents. We also acknowledge Roderic Schlotzhauer and Cheston Callais from the MU Physics Machine Shop for their design input and fabrication of our custom endoscopy platform and strategic modifications to commercial endoscopes and micromanipulators to meet our research needs. Our custom laryngeal motion tracking software was developed in collaboration with Dr. Filiz Bunyak and Dr. Ali Hamad (MU Electrical Engineering and Computer Science Department). We also thank Jim Marnatti from Karl Storz Endoscopy for providing guidance on otoscope selection. Finally, we would like to recognize numerous previous students/trainees in the Lever Lab whose contributions have informed the development of our current murine laryngoscopy protocol: Marlena Szewczyk, Cameron Hinkel, Abigail Rovnak, Bridget Hopewell, Leslie Shock, Ian Deninger, Chandler Haxton, Murphy Mastin, and Daniel Shu.
Atipamezole | Zoetis | Antisedan; 5 mg/mL | Parsippany-Troy Hills, NJ |
Bioamplifier | Warner Instrument Corp. | DP-304 | Hamden, CT |
Concentric EMG needle electrode | Chalgren Enterprises, Inc. | 231-025-24TP; 25 mm x 0.3 mm/30 G | Gilroy, CA |
Cotton tipped applicator (tapered) | Puritan Medical Products | REF 25-826 5W | Guilford, ME |
Data Acquisition System | ADInstruments | PowerLab 8/30 | Colorado Springs, CO |
DC Temperature Control System – for endoscopy platform | FHC, Inc. | 40-90-8D | Bowdoin, ME |
Electrophysiology recording software | ADInstruments | LabChart 8 with video capture module | Colorado Springs, CO |
Endoscope monitor | Karl Storz Endoscopy-America | Storz Tele Pack X monitor | El Segundo, CA |
Glycopyrrolate | Piramal Critical Care | NDC 66794-204-02; 0.2 mg/mL | Bethlehem, PA |
Ground electrode | Consolidated Neuro Supply, Inc. | 27 gauge stainless steel, #S43-438 | Loveland, OH |
Isoflurane induction chamber | Braintree Scientific, Inc. | Gas Anesthetizing Box – Red | Braintree, MA |
Ketamine hydrochloride | Covetrus North America | NDC 11695-0703-1, 100 mg/mL | Dublin, OH |
Metal spatula to decouple epiglottis and velum | Fine Science Tools | Item No. 10091-12; | Foster City, CA |
Micro-brush to remove food/secretions from oral cavity | Safeco Dental Supply | REF 285-0023, 1.5 mm | Buffalo Grove, IL |
Mouse-size heating pad for endoscopy platform | FHC, Inc. | 40-90-2-07 – 5 x 12.5 cm Heating Pad | Bowdoin, ME |
Ophthalmic ointment (sterile) | Allergan, Inc. | Refresh Lacri-lube | Irvine, CA |
Otoscope | Karl Storz | REF 1232AA | El Segundo, CA |
Pneumogram Sensor | BIOPAC Systems, Inc. | RX110 | Goleta, CA |
Pulse oximetry – Vetcorder Pro Veterinary Monitor | Sentier HC, LLC | Part No. 710-1750 | Waukesha, WI |
Rat-size heating pad for endoscopy platform | FHC, Inc. | 40-90-2 – 12.5X25cm Heating Pad | Bowdoin, ME |
Sterile needles for drug injections | Becton, Dickinson and Company | REF 305110, 26 G x 3/8 inch, PrecisionGlide | Franklin Lakes, NJ |
Sterile syringes for drug injections | Becton, Dickinson and Company | REF 309628; 1 mL, Luer-Lok tip | Franklin Lakes, NJ |
Surgical drape to cover induction cage for dark environment | Covidien LP | Argyle Surgical Drape Material, Single Ply | Minneapolis, MN |
Surgical tape to secure pneumograph sensor to abdomen | 3M Health Care | #1527-0, 1/2 inch | St. Paul, MN |
Transparent blanket for thermoregulation | The Glad Products Company | Press’n Seal Cling Film | Oakland, CA |
Video editing software | Pinnacle Systems, Inc. | Pinnacle Studio, v24 | Mountain View, CA |
Water circulating heating pad – for anesthesia induction/recovery station | Adroit Medical Systems | HTP-1500 Heat Therapy Pump | Loudon, TN |
Xylazine | Vet One | NDC 13985-701-10; Anased, 100 mg/mL | Boise, ID |
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