Minimally Invasive Murine Laryngoscopy for Close-Up Imaging of Laryngeal Motion During Breathing and Swallowing

Published: December 01, 2023

doi:

Teresa E. Lever², Ashley Kloepper, Lauren Welby, Megan Haney, Shakera Fudge, Chase Seiller, Sophia Kington, Brayton Ballenger, Nicole L. Nichols

¹Department of Otolaryngology – Head and Neck Surgery,University of Missouri School of Medicine, ²Department of Biomedical Sciences,University of Missouri College of Veterinary Medicine, ³Division of Otolaryngology – Head and Neck Surgery,Texas A&M Health Science Center College of Medicine – Baylor Scott & White, ⁴Department of Veterinary Pathobiology,University of Missouri College of Veterinary Medicine, ⁵Department of Medicine,University of Missouri School of Medicine

Summary

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.

Abstract

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.

Introduction

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.

Protocol

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 …

Representative Results

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 investi…

Discussion

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 t…

Disclosures

The authors have nothing to disclose.

Acknowledgements

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.

Materials

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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