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. Author manuscript; available in PMC: 2019 Oct 15.
Published in final edited form as: Neuroscience. 2018 Sep 1;390:303–316. doi: 10.1016/j.neuroscience.2018.08.026

Hypoglossal motor neuron death via intralingual CTB-saporin (CTB-SAP) injections mimic aspects of amyotrophic lateral sclerosis (ALS) related to dysphagia

Lori A Lind 1, Erika R Murphy 2, Teresa E Lever 1,2,3, Nicole L Nichols 1,4,*
PMCID: PMC6168367  NIHMSID: NIHMS1505736  PMID: 30179644

Abstract

Amyotrophic lateral sclerosis (ALS) is a devastating disease leading to degeneration of motor neurons and skeletal muscles, including those required for swallowing. Tongue weakness is one of the earliest signs of bulbar dysfunction in ALS, which is attributed to degeneration of motor neurons in the hypoglossal nucleus in the brainstem, the axons of which directly innervate the tongue. Despite its fundamental importance, dysphagia (difficulty swallowing) and strategies to preserve swallowing function have seldom been studied in ALS models. It is difficult to study dysphagia in ALS models since the amount and rate at which hypoglossal motor neuron death occurs cannot be controlled, and degeneration is not limited to the hypoglossal nucleus. Here, we report a novel experimental model using intralingual injections of cholera toxin B conjugated to saporin (CTB-SAP) to study the impact of only hypoglossal motor neuron death without the many complications that are present in ALS models. Hypoglossal motor neuron survival, swallowing function, and hypoglossal motor output were assessed in Sprague Dawley rats after intralingual injection of either CTB-SAP (25 g) or unconjugated CTB and SAP (controls) into the genioglossus muscle. CTB-SAP treated rats exhibited significant (p≤0.05) deficits vs. controls in: 1) lick rate (6.0±0.1 vs. 6.6±0.1 Hz; 2); hypoglossal motor output (0.3±0.05 vs. 0.6±0.10 mV); and 3) hypoglossal motor neuron survival (398±34 vs. 1018±41 neurons). Thus, this novel, inducible model of hypoglossal motor neuron death mimics the dysphagia phenotype that is observed in ALS rodent models, and will allow us to study strategies to preserve swallowing function.

Keywords: swallowing, neurodegenerative disease, videofluoroscopy, motor output

Introduction

Amyotrophic lateral sclerosis (ALS) is a progressive neurodegenerative disorder in which the loss of both upper and lower motor neurons leads to weakness and paralysis of skeletal muscles (Kiernan et al., 2011). The most serious consequences are the result of neuronal death in the phrenic and hypoglossal motor nuclei. Respiratory failure is the most common cause of death and is often due to diaphragmatic weakness resulting from the loss of phrenic motor neurons (Corcia et al., 2008; Kurian et al., 2009; Kiernan et al., 2011). The loss of hypoglossal motor neurons results in impairments in speech (Langmore and Lehman, 1994; Tomik and Guiloff, 2010), swallowing (Kawai et al., 2003; Higo et al., 2004; Jani and Gore, 2016; Onesti et al., 2017), and the ability to maintain upper airway patency (Perrin et al., 2004). Dysphagia (swallowing dysfunction) often leads to malnutrition and aspiration pneumonia which may in turn result in respiratory failure and death (Hadjikoutis et al., 2001). Despite the fundamental importance of maintaining respiratory and swallowing function in ALS patients, research in these areas has been limited and the only treatments currently available are strictly palliative.

We hypothesize that patient survival and quality of life can be improved by increasing the plasticity of surviving phrenic and hypoglossal motor neurons, thereby allowing them to maintain motor output and preserve respiratory and swallowing function for longer periods of time. Testing this hypothesis requires a translational animal model to study methods of inducing plasticity and discerning the mechanisms by which this plasticity occurs. The main rodent model of dysphagia in ALS is a transgenic mouse model (SOD1G93A; Gurney et al., 1994) that develops progressive dysphagia characterized by decreased lick and swallow rates relative to healthy controls (Lever et al., 2009; 2010; 2015a). These symptoms of dysphagia are attributed to degeneration of several brainstem nuclei involved in swallowing, including the hypoglossal nucleus (Lever et al., 2009; 2010). However, the degree and rate of neuronal death cannot be controlled in this mouse model, and degeneration is not limited to the hypoglossal motor neurons. Thus, we have developed an inducible model in which we used intralingual injections of cholera toxin B conjugated to saporin (CTB-SAP) into the genioglossus muscle to produce targeted death of hypoglossal motor neurons in the brainstem medulla.

CTB-SAP consists of a tracer molecule (CTB) linked to a toxin (SAP) via a disulfide bond (Llewellyn-Smith et al., 2000). Following intralingual injection, CTB-SAP is taken up by nearby axons and then retrogradely transported to the neuron cell body (Llewellyn-Smith et al., 1999; 2000; Lujan et al., 2010). The CTB component then binds to GM1 (Galactosyl-N-Acetylgalactosaminyl) receptors on the cell membrane of motor neurons (Lian and Ho, 1997), allowing the entire CTB-SAP construct to enter the cell through endocytosis (Lencer and Tsai, 2003), where CTB and SAP will then dissociate (Llewellyn-Smith et al., 2000). SAP is a ribosomal inactivating protein and once released from the CTB it binds to ribosomes and renders them incapable of protein synthesis, resulting in the apoptotic death of the neuron within hours to days (Llewellyn-Smith et al., 1999; Lujan et al., 2010).

We previously demonstrated that CTB-SAP injected into the intrapleural space of rats is taken up by nearby axons of phrenic motor neurons and selectively causes death of phrenic motor neurons at a level that recapitulates what is seen in the SOD1G93A rat model of ALS (Nichols et al., 2015b). Thus, we hypothesized that injection of CTB-SAP directly into the tongue would result in the targeted cell death of hypoglossal motor neurons while sparing all other cells, and thereby result in decreased hypoglossal output and deficits in swallowing function. We found that CTB-SAP treated rats did exhibit significant deficits when compared to controls including decreased hypoglossal motor neuron survival, hypoglossal motor output, lick and swallow rates. This suggests that CTB-SAP-induced hypoglossal motor neuron death did result in hypoglossal and swallowing deficits, and mimics the dysphagia phenotype that is observed in ALS rodent models.

Experimental Procedures

Animals

Experiments were conducted on 22 adult (3-4 months) male Sprague Dawley rats (Envigo Colony 208; Indianapolis, IN). Animals were housed in pairs and maintained under a 12:12 light:dark cycle. A standard commercial pelleted diet and water were offered ad libitum except where otherwise noted. All procedures involving animals were approved by the Institutional Animal Care and Use Committee at the University of Missouri in accordance with National Institutes of Health Guide for the Care and Use of Laboratory Animals. The University of Missouri is an AAALAC-accredited institution that operates under Animal Welfare Assurance ID A3394-01.

Intralingual injections

All rats received intralingual injections into the genioglossus muscle to selectively target hypoglossal motor neurons as previously described (Lee et al., 2011; ElMallah et al., 2012). Animals in the treatment group received 25 μg of CTB-SAP (dissolved in phosphate buffered saline (PBS; n=11); Advanced Targeting Systems, San Diego, CA) to selectively cause death of hypoglossal motor neurons, plus extra CTB (25 μg dissolved in doubly distilled water, Calbiochem, Billerica, MA) to label surviving hypoglossal motor neurons. Control treated animals received CTB (25 μg; n=11) unconjugated to saporin (25 μg; dissolved in PBS; Advanced Targeting Systems, San Diego, CA) to show that neither one alone causes hypoglossal motor neuron death.

Rats received intralingual injections while under isoflurane anesthesia (2-3%) via a nose cone (Fig. 1). Specifically, rats were placed on a custom tilt table in a supine position, and the head was gently immobilized using ear bars (Fig. 1). The mouth was gently held open by a custom string pulley system looped around the mandibular incisors (Fig. 1), and fine forceps were used to carefully lift the tongue to visualize the frenulum connecting the underside of the tongue to the floor of the mouth. A 26 gauge needle was inserted into the midline frenulum at a 45° angle (Fig. 1) to target the genioglossus muscle within the tongue base. Half of the injection was administered while the needle was at maximum depth (8 mm), and the other half was given after the needle was retracted halfway (4 mm). In a subset of rats (6 rats per group), two 0.5 mm tantalum beads (Bat-tec, Los Angeles, CA) were injected into the tip of the tongue via a 21 gauge × 1.5 inch needle and custom-made metal stylus to enable visualization of the tongue during radiographic swallowing function studies. Rats were then monitored for overt signs of feeding and drinking behavior, as described below.

Figure 1: Intralingual injection setup.

Figure 1:

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A. A customized tilt table (1) is used for intralingual injections. The setup includes ears bars (2) to immobilize the head, nose cone (3) and anesthesia (4) / vacuum scavenging (5) lines for isoflurane anesthesia, a jaw pulley system (6) including incisor loop (7) and 30 g weight (8) to gently hold the mouth open, and a mini gooseneck light (9) for intraoral visualization during intralingual injection. B. An anesthetized rat on our customized setup receiving a CTB-SAP injection (10) via a 26 gauge needle at an approximate 45° angle. The incisor loop (7) of our jaw pulley system is positioned at the base of the mandibular incisors to gently hold the mouth open during intralingual injection.

Videofluoroscopic swallow study

The videofluoroscopic swallow study (VFSS) was conducted on all rats prior to tongue injections (i.e., baseline), and on day 8 following injections. We also conducted VFSS on day 4 following tongue injections on the first half of rats studied to give us feedback on whether we would have to change our methodology. We discontinued VFSS studies at the 4 day time-point since we did not detect a significant functional deficit, and thus we are only reporting VFSS data from our baseline and day 8 time-points. Rats were water restricted overnight for ~14 hours prior to VFSS testing to increase their willingness to drink while in the fluoroscope. During the water restriction, a custom-built polycarbonate test chamber was placed in the home cage for acclimation. The test chambers (10 cm high × 7.5 cm wide × 25 cm long) were modelled after those previously described for mice (Lever et al., 2015a), and contained ventilation holes in the ceiling on one end and 2 removable end caps (one of which contained an attached peg bowl). The next day, rats underwent VFSS testing in the same test chamber that was in the home cage overnight. After placing the rat in the test chamber, it was transferred to a custom-designed miniature fluoroscope for VFSS testing in the lateral plane, following a protocol previously developed for mice (Lever et al., 2015a). In short, our fluoroscope (Glenbrook Technologies, Randolph, NJ) is a low energy (40 kilovolts and 0.2 milliamperes) system that uses a custom, motorized lift table to aid in remote positioning of the rat’s head within the fluoroscope beam during VFSS testing. An iohexol solution (25% iohexol added to vanilla-flavored sugar water; 30% sucrose in water containing 120 microliters of non-alcoholic vanilla extract (per 150 ml of sugar water)) was delivered into the peg bowl within the test chamber via a syringe delivery system consisting of a 10 mL syringe and ~1 meter of polyethylene tubing (PE240). A webcam was focused on the chamber and projected onto a computer monitor to enable continuous visualization of the rats’ movement. To minimize radiation exposure, the x-ray beam was manually activated via a foot pedal only when the rat was actively drinking from the peg bowl. Since our fluoroscope was initially designed for mice, the field of view (5 cm diameter) was too small to encompass the entire region of interest in one position for rats. Thus, we captured videos (30 frames/second) at two positions for each rat: 1) Position 1 focused on the water bowl and was used to observe the entire jaw and tongue during drinking, which allowed us to calculate lick rate and assess tongue motility (see below); and 2) Position 2 was focused on the vallecular space between the tongue base and epiglottis, which allowed us to calculate swallowing rate by visualizing the accumulation of fluid in the vallecular space and its transfer to the esophagus with each swallowing event. All videos were blindly analyzed independently by two trained reviewers, and any discrepancies in the two analyses were further examined by additional blinded reviewers until consensus was reached. Videos were analyzed frame by frame using Pinnacle Studio 14 software (Pinnacle Systems Inc., Mountain View, CA). One jaw cycle was defined as the jaw moving from maximally open to closed to maximally open again. This measurement is used to represent lick rate because the tongue is not always visible on fluoroscopy, and maximum jaw opening corresponds to maximum tongue protrusion (Lever et al., 2015a). Five episodes of uninterrupted drinking lasting 2 seconds (60 frames) each were analyzed. Lick and swallow rates were calculated for each of the 5 episodes and then averaged for statistical analysis.

In vivo neurophysiology

Experimental preparation.

Nine days following intralingual injections, rats underwent in vivo neurophysiology procedures as described previously (Hoffman et al., 2012; Nichols et al., 2012, 2015b). Rats were anesthetized via chamber induction with isoflurane, and then maintained with 3.5% isoflurane in 50% O2 (balance N2). All rats were tracheotomized and ventilated (Small Animal SAR-1000 Ventilator; CWE, Ardmore, PA, USA; tidal volume ~2.5 ml, frequency ~70). Bilateral, mid-cervical vagotomies were performed to prevent entrainment to the ventilator. A polyethylene catheter (PE50 ID: 0.58 mm, OD: 0.965 mm; Intramedic, MD) was inserted into the right femoral artery to monitor blood pressure (APT300 Pressure Transducer, Harvard Apparatus, Holliston, MA, USA) and blood gases (ABL80 Flex, Radiometer, Brea, CA). A second catheter was placed in the femoral vein to allow administration of a continuous infusion of a 1:2:0.13 mixture of 6% hetastarch in 0.9% sodium chloride, lactated Ringer’s solution, and 8.4% sodium bicarbonate at a rate of 1.5-4 ml/kg/hr in order to maintain blood volume, fluid, and acid-base balance. Body temperature was monitored using a rectal thermistor probe (Physitemp, Clifton, NJ, USA) and maintained at 37.5±1 C by means of a custom-made heated surgical table. End-tidal PCO2 (PETCO2)) was maintained at ~45 mmHg as measured by a flow-through carbon dioxide analyzer with a sufficient response time to measure PETCO2 in rats (CapStar-100, CWE, Ardmore, PA). The left phrenic and hypoglossal nerves were isolated (dorsal approach), cut distally, desheathed, and covered with a saline-soaked cotton ball until the start of the experimental protocols. Here, we isolated and recorded (see below) from the phrenic nerve as an internal control to show that motor neuron death and deficits were limited to the hypoglossal nucleus. Once the surgical procedures were completed, rats were converted to urethane anesthesia (1.85 g/kg administered intravenously over 20-30 minutes). The adequacy of anesthesia was tested before protocols commenced, and immediately after the protocol was complete; adequacy of anesthetic depth was assessed as the lack of pressor or respiratory neural response to a toe pinch with a hemostat (Bach and Mitchell, 1996; Hoffman et al., 2012; Nichols et al., 2012; Nichols et al., 2015b). Once rats were converted to urethane, a minimum of 1 h was allowed before experiments commenced.

Nerve recordings.

The previously isolated left phrenic and hypoglossal nerves were submerged in mineral oil and placed on bipolar silver electrodes to record nerve activity. Neural signals were amplified (10,000X), band-pass filtered (300-10,000 Hz, Model 1800, A-M Systems, Sequim, WA), and full-wave rectified and integrated (50 ms time constant, MA-821, CWE Inc., Ardmore, PA, USA). Integrative nerve bursts were digitized (8 kHz) and analyzed using WINDAQ data acquisition system (DATAQ Instruments, Akron, OH). Pancuronium bromide (2.5 mg/kg) was administered intravenously for neuromuscular blockade to prevent spontaneous efforts to breathe (Bach and Mitchell, 1996).

To begin the protocols, the apneic CO2 threshold was determined by lowering PETCO2 until nerve activity ceased for at least one minute. The PETCO2 was then slowly increased until the recruitment threshold was reached (i.e. nerve activity resumed; Bach and Mitchell, 1996). PETCO2 was then raised 2 mmHg above the recruitment threshold, and ~15-20 min were allowed to establish stable baseline activity. Blood samples were collected to measure partial arterial pressures of oxygen (PaO2) and carbon dioxide (PaCO2), and were measured during baseline, the two hypercapnic challenges, and the hypercapnic + hypoxic episode. PaO2 was ≥150 mm Hg during baseline and during hypercapnic challenges, but was between 35–45 mm Hg during hypercapnia + hypoxia. First, the PaCO2 was increased by 20 mmHg above baseline for 5 minutes followed by another 5 minutes at 40 mmHg above baseline. Finally, the PaCO2 levels were maintained at 40 mmHg above baseline while the PaO2 levels were decreased to 35-45 mmHg for 5 minutes before returning all gases to baseline levels at the end of the experiment.

Immunohistochemistry

Immediately after the in vivo neurophysiology experiments were complete on day 9, rats were transcardially perfused with 4% paraformaldehyde in 0.1M phosphate buffer saline (PBS, pH~7.4). The spinal cord, brain, and brainstem were harvested and post-fixed with 4% paraformaldehyde overnight, and then cryoprotected in graded sucrose (20 and 30%) at 4 °C until sinking. The midbrain, pons, medulla, and cervical spinal cord were then transversely cut into 40 μm thick sections using a freezing sliding microtome (Leica SM 2010R, Germany) and stored in antifreeze (30% ethylene glycol, 30% glycerol and 1X PBS) at −20 °C until processed for motor neuron counting.

Eight sections of the medulla were selected that were representative of the entire hypoglossal nucleus of each rat (i.e., 8 sections were selected that were evenly distributed throughout the entire length of the hypoglossal nucleus). Six sections of the 4th cervical spinal cord segment containing the phrenic motor nucleus were included as an internal control. For 16 of the animals (8 controls and 8 CTB-SAP treated), additional sections were collected from other parts of the medulla, pons, and midbrain to study possible off-target staining. Sections were first separated and washed with 1X PBS three times for five minutes; each rat’s tissue was contained in a separate well for each region of interest. To prevent non-specific antibody binding, blocking solution consisting of 5% normal donkey serum (NDS), 1X PBS, and 0.2% Triton was added to each well containing tissue sections and incubated for 1 hour at room temperature. Primary antibody solution was added, consisting of 5% NDS, 1X PBS, 0.1% Triton, and the antibody against cholera toxin B subunit (CTB; goat polyclonal, 1:2000, Calbiochem; Billerica, MA). Sections were incubated overnight in the primary antibody solution on a shaker at 4 C. The following day, tissues were washed three times with 1X PBS (five minutes each), and then incubated in the secondary antibody solution composed of 5% NDS, 1X PBS, 0.1% Triton, and secondary antibody (donkey anti-goat Alexa-Fluor 555, 1:1000; Molecular Probes, Eugene, OR) on a shaker for two hours in the dark at room temperature. Following incubation, tissues were washed with 1X PBS using the same procedure (3 × 5min). Sections were then mounted on positively charged glass slides, covered with ProLong® Gold Antifade Reagent (Molecular Probes, Eugene, OR) to prevent quenching of fluorescence, cover-slipped, and air-dried in the dark. Slides were stored at −20 C until quantification of staining was performed using an epifluorescence microscope (Leica DM4000; Germany) at 20x magnification. Sections incubated without primary or secondary antibodies served as negative controls. In addition, to determine the ratio of total hypoglossal motor neurons labeled by CTB, we performed identical immunohistochemistry experiments as explained above but with both anti-CTB (donkey anti-goat Alexa-Fluor 555 as the secondary) and the antibody against neuronal nuclei (NeuN; mouse monoclonal, 1:500, Millipore; Burlington, MA; donkey anti-mouse Alexa Fluor 488 (1:1000) as the secondary) on tissue from a small cohort of the control rats (n=3).

Motor Neuron Counts

Cells were manually counted at 20X using an epifluorescence microscope (Leica DM4000; Germany) by a blinded investigator. Stereo Investigator software (MBF Bioscience, Williston, VT) was used to outline each region of interest, mark counted cells, and save images for future reference. For each section, the hypoglossal nucleus was divided into left and right halves and dorsal and ventral compartments based on Aldes (1995), McClung and Goldberg (1999), and Behan et al. (2012), and the counts from each quadrant were recorded separately. Surviving motor neurons were characterized as those exhibiting CTB(+) staining, and an identifiable cell body and nucleus. A Grubb’s outlier test (GraphPad software) was performed and the most rostral and caudal sections of the hypoglossal nucleus consistently contained no CTB(+) neurons in either controls or CTB-SAP treated animals; therefore, these sections were excluded from the final analysis, leaving 6 sections per animal for quantification. These six sections covered the range of the hypoglossal nucleus extending from approximately Bregma −12.96 mm to −14.40 mm for a length of 1440 m (Paxinos and Watson, 2005). The hypoglossal motor neuron counts were then extrapolated to estimate the number of neurons in the sampled region for each animal (1440 m for length of the hypoglossal nucleus; 40 m sections). Similar extrapolations were made for all other nuclei in which we found off-target staining of CTB(+) cells: trigeminal motor nucleus (~840 μm length; 40 m sections), facial motor nucleus (~1320 μm length; 40 m sections), and salivatory nucleus (~1920 μm length; 40 m sections).

Statistical analysis

VFSS and integrated nerve burst amplitude data were compared with a two-way repeated measures ANOVA, with treatment group (control vs. CTB-SAP) and time-point or level (i.e., baseline and end-point for VFSS time-points; baseline and hypercapnia+hypoxia (maximum chemoreceptor stimulation) for nerve amplitude levels) as factors. Integrated nerve burst amplitudes were averaged over 1 min during baseline, during the two hypercapnic challenges, and during the hypercapnia+hypoxia challenge. Only baseline and hypercapnia+hypoxia data are reported for integrated nerve burst amplitudes, where nerve burst amplitude is reported as the voltage of the integrated signal. For histological analysis, motor neuron survival was averaged across sections within animals and then a t-test was used to compare across control and CTB–SAP treatment groups. We tested for and found normal distributions and equal variances in our sample distributions. If significant differences were indicated, multiple comparisons were made using a Fisher's least significant difference (LSD) post hoc test (Sigma Plot version 13.0; Systat Software Inc., San Jose, CA, USA). All differences between groups were considered significant if p≤0.05; all values are expressed as means ± 1 S.E.M. Multiple linear regression analyses were performed between baseline or maximal hypoglossal activity with ventral hypoglossal motor neuron survival, lick rate and swallow rate with ventral hypoglossal motor neuron survival, and lick rate and swallow rate with baseline or maximal hypoglossal activity.

Results

Hypoglossal motor neurons are targeted by intralingual injections of CTB-SAP

Intralingual injections of CTB-SAP into the genioglossus muscle selectively targeted motor neurons whose axons terminate in the tongue. Motor output to the tongue is supplied by the hypoglossal nucleus, with the ventral compartment supplying the protrusor muscles of the tongue (including the genioglossus), and the dorsal compartment supplying the retrusor muscles (Aldes, 1995; McClung and Goldberg, 1999; Behan et al., 2012). Figure 2 includes representative photomicrographs of the hypoglossal nucleus from both a control rat and a CTB-SAP treated rat at day 9 showing how the compartments were divided for neuronal counts. The CTB-SAP treated rats had significantly fewer CTB(+) neurons relative to controls in both the dorsal (157±26 vs. 312±29; t20 = 3.992, p < 0.001 and ventral (398±34 vs.1018±41; t20 = 11.634, p < 0.001) compartments (Fig. 2). No CTB(+) neurons were observed in the medullary tissue directly surrounding the hypoglossal nucleus or in any of the spinal cord segments containing the phrenic motor nucleus (data not shown). To determine the ratio of hypoglossal motor neurons labeled by CTB and how many hypoglossal motor neurons were targeted by CTB-SAP, we performed co-labeling experiments with anti-CTB and anti-NeuN in three control rats. We found that CTB labels 66% of the total NeuN labeled ventral hypoglossal motor neurons, and CTB labels 15% of the total NeuN labeled dorsal hypoglossal motor neurons (data not shown). In this study, CTB-SAP treated rats have 39% ventral hypoglossal motor neuron survival (i.e., 61% death) and 50% dorsal hypoglossal motor neuron survival (i.e., 50% death) (Fig. 2). Thus, we targeted 40% of the total ventral hypoglossal motor neurons and 7.5% of the total dorsal hypoglossal motor neurons. Our goal in this study was to target approximately half of the ventral hypoglossal motor neurons (motor neurons responsible for tongue protrusion) with our CTB-SAP genioglossal injections; thus, we have successfully achieved this goal by targeting 40% of the total ventral hypoglossal motor neurons.

Figure 2: Hypoglossal motor neuron survival in CTB-SAP treated rats.

Figure 2:

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A-F. Photomicrographs (4X A,D; 20X B,C,E,F) of representative CTB stained sections from the brainstem medulla of a control (A-C) and CTB-SAP treated rat (D-F) at day 9. In A and D, the yellow outline (top) indicates the dorsal (retrusor) compartment of the hypoglossal nucleus and the teal outline (bottom) indicates the ventral (protrusor) compartment. The dashed white borders outline the regions shown at 20X in B, C, E, and F. Note that the neurons in B and C all look healthy while many of the neurons in E and F (particularly in the bottom half of each image) are more spherical and lack visible processes and nuclei (yellow arrows indicate dead/dying neurons which were excluded from the hypoglossal motor neuron survival counts). G-H. Hypoglossal motor neuron survival in the dorsal (G) and ventral (H) compartments. CTB-SAP treated rats (gray bars) had significantly reduced numbers of surviving CTB(+) neurons compared to control rats (black bars) for both compartments (*; p≤0.05). Means ± 1 SEM. Scale bar at 4X = 200 μm, 20X = 50 μm.

Hypoglossal, not phrenic, motor output is reduced in rats treated with intralingual injections of CTB-SAP

Hypoglossal motor output (recorded 9 days post injection) from controls and CTB-SAP treated rats during baseline and graded chemoreceptor stimulation are illustrated in Figure 3. Representative compressed, integrated hypoglossal neurograms from a control rat and a CTB-SAP treated rat are depicted in Figure 3A. A two-way repeated measures ANOVA revealed significant differences for hypoglossal motor output for level (F1(between groups),20(within groups) = 50.998, p < 0.001), where both treatment groups (controls and CTB-SAP treated rats) responded to chemoreceptor challenge with increased hypoglossal motor output (Fisher’s LSD post hoc = p < 0.001 for both groups; Fig. 3B). In addition, a two-way repeated measures ANOVA revealed significant differences for hypoglossal motor output for treatment group (F1,20 = 9.103, p = 0.008), where CTB-SAP treated rats had a significantly lower output at both baseline and maximum chemoreceptor stimulation (0.3±0.05 mV and 0.9±0.15 mV, respectively) compared to controls (0.6±0.11 mV and 1.6±0.24 mV, respectively; Fisher’s LSD post hoc = p = 0.004 for baseline and p = 0.003 for maximum chemoreceptor stimulation; Fig. 3B).

Figure 3: Hypoglossal motor output in control and CTB-SAP treated rats.

Figure 3:

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A. Representative traces of compressed integrated hypoglossal neurograms in control and CTB-SAP treated rats recorded 9 days after intralingual injection. Spontaneous hypoglossal activity was recorded at baseline and during graded levels of hypercapnia (5 minutes each with inspired CO2 levels that were 20 mmHg and 40 mmHg above baseline), followed by hypercapnia+hypoxia (inspired CO2 at 40 mmHg above baseline and O2 decreased from 21% to 11%). B. Comparison of integrated hypoglossal amplitude at baseline and in response to maximum chemoreceptor stimulation (hypercapnia+hypoxia). Both groups showed increased hypoglossal motor output in response to hypercapnia+hypoxia (#; p≤0.05), but CTB-SAP treated rats (gray line with gray circles) had decreased output relative to controls (black line with black triangles) at both time points (*; p≤0.05). Means ± 1 SEM.

We also simultaneously studied phrenic motor output as an internal control from CTB-SAP treated rats and controls since phrenic motor neurons (found in the cervical 3-6 segments of the spinal cord) should not be targeted by the intralingual CTB-SAP injection. Representative compressed, integrated phrenic neurograms from a control rat and a CTB-SAP treated rat are depicted in Figure 4A. Phrenic motor output was not affected by intralingual injections of CTB-SAP at baseline or chemoreceptor challenge when compared to controls (F1,20 = 0.174, p = 0.681; Fig. 4). As expected, both controls and CTB-SAP treated rats responded to chemoreceptor challenge with increased phrenic motor output (controls: 0.9±0.23 mV at baseline and 2.0±0.45 mV during maximum chemoreceptor stimulation; CTB-SAP: 1.0±0.22 mV at baseline and 2.2±0.44 mV during maximum chemoreceptor stimulation, F1,20 = 38.183, p < 0.001; Fisher’s LSD post hoc = p < 0.001 for both treatment groups; Fig. 4B).

Figure 4: Phrenic motor output in control and CTB-SAP treated rats.

Figure 4:

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A. Representative traces of compressed integrated phrenic neurograms in control and CTB-SAP treated rats recorded 9 days after intralingual injection. Spontaneous phrenic activity was recorded at baseline and during graded levels of hypercapnia (5 minutes each with inspired CO2 levels that were 20 mmHg and 40 mmHg above baseline), followed by hypercapnia+hypoxia (inspired CO2 at 40 mmHg above baseline and O2 decreased from 21% to 11%). B. Comparison of integrated phrenic amplitude at baseline and in response to maximum chemoreceptor stimulation (hypercapnia+hypoxia). Both groups showed increased phrenic output in response to hypercapnia+hypoxia (#; p≤0.05), but there was no difference between the two groups (controls = black line with black triangles; CTB-SAP = gray line with gray circles; p>0.05). Means ± 1 SEM.

Intralingual injections of CTB-SAP result in dysphagia and compensatory swallowing behaviors

Eight days after intralingual injections, we observed a number of swallowing-related behavioral changes in CTB-SAP treated rats (Fig. 5), similar to what has been observed in SOD1G93A mice and ALS patients (Kühnlein et al., 2008; Jani and Gore, 2016; Onesti et al, 2017; T.E. Lever, unpublished observations). For example, CTB-SAP treated rats held their head in a more vertical position with the nose pointed down (Fig. 5), similar to what is observed in ALS patients with head drop (Gourie-Devi et al., 2003). Also, as indicated by the position of the tongue-implanted beads during active drinking in Figure 5, the tip of the tongue appears to be pulled further back into the mouth during maximum tongue retraction, and the tongue failed to consistently contact the liquid in the bowl during maximum tongue protrusion. In addition, CTB-SAP treated rats frequently placed a forepaw on the edge of the bowl for added stability while drinking (Fig. 5), similar to what is observed in SOD1G93A mice (T.E. Lever, unpublished observations). A two-way repeated measures ANOVA revealed significant differences in swallowing function for treatment group (F1,20 = 11.997, p = 0.002 for lick rate; F1,20 = 5.967, p = 0.026 for swallow rate) and time-point (F1,20 = 15.983, p < 0.001 for lick rate; F1,20 = 5.091, p = 0.038 for swallow rate). Fisher’s LSD post hoc tests showed that controls and CTB-SAP treated rats had similar swallowing function at baseline (p = 0.071 for lick rate; p = 0.184 for swallow rate; Fig. 6). However, CTB-SAP treated rats showed a significant decrease in swallowing function at the study end-point compared to controls, which included slower lick rate (6.0±0.1 vs. 6.6±0.1 Hz, respectively; p = 0.001) and swallow rate (2.0±0.1 vs. 2.5±0.2 Hz, respectively; p = 0.026) (Fig. 6). CTB-SAP treated rats also showed a significant decrease in swallowing function at end-point compared to their own baseline (p = 0.001 for lick rate and p = 0.044 for swallow rate; Fig. 6). Thus, swallowing deficits and compensatory behavioral changes observed in CTB-SAP treated rats are consistent with what is observed in SOD1G93A mice and ALS patients (Kawai et al., 2003; Higo et al., 2004; Lever et al., 2009; 2010; 2015a; Jani and Gore, 2016).

Figure 5: Single frames taken from videoradiographs of a rat prior to intralingual CTB-SAP treatment (A and B) and at the end-point of the study (8 days post injection; C and D).

Figure 5:

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Images were selected to illustrate maximum tongue protrusion (A and C) and maximum tongue retraction (B and D). Note the following changes that occurred after injection of CTB-SAP: 1) Initially the tongue extended all the way to the meniscus of the flavored water (denoted by white arrow), but after intralingual CTB-SAP injection, the tongue failed to extend beyond the incisors at maximum protrusion; 2) The tip of the tongue also moved further back in the mouth during maximum retraction (black arrowheads indicate 2 tantalum beads placed in the tip of the tongue to make it more readily visible during fluoroscopy); 3) The head was held in a more vertical position (i.e. the chin was tucked); and 4) A paw was placed on the edge of the bowl for added stability.

Figure 6: Comparison of swallowing metrics in control and CTB-SAP treated rats.

Figure 6:

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Comparison of swallow rate (A; number of swallows per 2 second interval) and lick rate (B; number of maximum jaw openings in 1 second) in controls (black bars) vs. CTB-SAP treated rats (gray bars) at baseline and end-point. The CTB-SAP treated rats showed deficits in both lick and swallow rates at 8 days post injection relative to both the control rats (*; p≤0.05) and their own baseline function (#; p≤0.05).

Correlations between hypoglossal motor output, ventral hypoglossal motor neuron survival and swallowing behavior

Multiple regression analyses were performed, including baseline or maximal (hypercapnia + hypoxia) hypoglossal output with hypoglossal motor neuron survival (Figs. 7A & B); swallow rate and lick rate with hypoglossal motor neuron survival (Figs. 7C & D); swallow rate with baseline and maximal hypoglossal output (Figs. 7E & F); and lick rate with baseline and maximal hypoglossal output (Figs. 7G & H) for all rats in this study (n=22). Significant positive correlations were detected between: 1) maximal hypoglossal output with hypoglossal motor neuron survival (slope = 0.67; R2 = 0.20; F = 4.838, p = 0.04; Fig. 7B); 2) lick rate with hypoglossal motor neuron survival (slope = 5.7; R2 = 0.43; F = 14.751; p = 0.001; Fig. 7D); and 3) swallow rate with maximal hypoglossal output (slope = 1.7; R2 = 0.30; F = 8.459; p = 0.009; Fig. 7F). Significant correlations were not detected between baseline hypoglossal output with hypoglossal motor neuron survival (slope = 0.23; R2 = 0.13; F = 2.880; p = 0.105; Fig. 7A), swallow rate with hypoglossal motor neuron survival (slope = 1.8; R2 = 0.18; F = 4.245; p = 0.053; Fig. 7C), swallow rate with baseline hypoglossal output (slope = 2.1; R2 = 0.07; F = 1.445; p = 0.243; Fig. 7E), or lick rate with baseline or maximal hypoglossal output (baseline: slope = 6.1; R2 = 0.06; F = 1.173; p = 0.292; Fig. 7G; maximum: slope = 5.9; R2 = 0.17; F = 4.086; p = 0.057; Fig. 7H). Thus, hypoglossal motor neuron survival predicts maximal hypoglossal motor output and lick rate, and maximal hypoglossal output predicts swallow rate.

Figure 7: Multiple regression analyses between hypoglossal output and swallowing behavior with ventral hypoglossal motor neuron survival or swallowing behavior with hypoglossal output.

Figure 7:

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Multiple regression analyses between baseline or maximal hypoglossal output with ventral hypoglossal motor neuron survival (A, B), lick rate and swallow rate with ventral hypoglossal motor neuron survival (C, D), swallow rate with baseline or maximal hypoglossal output (E, F), and lick rate with baseline or maximal hypoglossal output (G, H). Significant correlations exist between maximal hypoglossal output with ventral hypoglossal motor neuron survival (p = 0.04), lick rate with ventral hypoglossal motor neuron survival (p = 0.001), and swallow rate with maximal hypoglossal output (p = 0.009). Thus, ventral hypoglossal motor neuron survival predicts maximal hypoglossal motor output and lick rate, and maximal hypoglossal motor output predicts swallow rate.

Non-targeted cell death was minimal following intralingual CTB-SAP injections

Since cell death is possible in regions outside of the hypoglossal nucleus following intralingual injections, tissue sections from the rostral medulla, pons, and midbrain of a subset of rats (8 controls and 8 CTB-SAP treated animals at day 9) were examined to look for CTB(+) cells. All animals had CTB(+) neurons in the trigeminal motor nucleus, facial nucleus, and superior salivatory nucleus. However, the number of CTB(+) cells was minimal and not significantly different between controls and CTB-SAP treated rats for the facial nucleus (t14 = 0.673, p = 0.538) and superior salivatory nucleus (t14 = 0.0869, p = 0.935; data not shown), but was significantly different between groups for the trigeminal motor nucleus (~40% loss in CTB-SAP treated rats; t14 = 2.603, p = 0.021 ; Fig. 8).

Figure 8: Trigeminal motor neuron survival in CTB-SAP treated rats.

Figure 8:

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A-F. Photomicrographs (larger images at 4X; insets at 20X) of representative CTB(+) stained sections from the region containing the trigeminal motor nucleus from a control (A-C) and CTB-SAP treated rat (D-F) at day 9. G. Trigeminal motor neuron survival. CTB-SAP treated rats (gray bars) had significantly reduced numbers of surviving CTB(+) neurons compared to control rats (black bars) (*; p≤0.05). Means ± 1 SEM. Scale bar at 4X = 400 μm, 20X = 50 μm.

Discussion

The main findings of this study are that intralingual injections of CTB-SAP into the genioglossus muscle at midline in rats results in: 1) targeted hypoglossal motor neuron death (Fig. 2); 2) decreased hypoglossal motor output (Fig. 3); 3) and swallowing deficits (Fig. 6). Thus, our data support our original hypotheses. However, we also identified additional important findings, including compensatory behavioral changes (Fig. 5) and off-target trigeminal motor neuron death (Fig. 8). Although these results were unanticipated, they are typically found in ALS patients. Specifically, the compensatory behaviors we observed (i.e., vertical head position) are similar to what is observed in ALS patients (i.e., dropped-head syndrome; Gourie-Devi et al., 2003). Furthermore, we also observed significant trigeminal motor neuron death, which is similar to what is observed in ALS patients (Lawyer and Netsky, 1953; Bonduelle, 1975; Hughes, 1982; DePaul et al., 1988) as well as the SOD1G93A mouse model (Gourie-Devi et al., 2003; Robbins et al., 2015). Thus, these findings strengthen the translational potential of our model for use in ALS research. Together our studies are consistent with intralingual CTB-SAP injections resulting in hypoglossal motor deficits and dysphagia.

Intralingual injections of CTB-SAP result in dysphagia

The deficits in lick and swallow rates exhibited by the CTB-SAP treated rats (Fig. 6) are very similar to those previously described for the SOD1G93A mouse model of ALS (Lever et al., 2009; 2010; 2015a). Some of the behavioral changes seen in the transgenic mouse model are also replicated in our inducible model. For example, the rats and mice both placed a forepaw on the edge of the bowl when drinking. There are some differences histologically, however. SOD1G93A mice show severe vacuolar degeneration in the mitochondria of the neuropil in parts of the brainstem, including the hypoglossal and trigeminal motor nuclei (Wong et al., 1995; Kong and Xu, 1998; Lever et al., 2009). We performed hematoxylin and eosin staining on 10 μm sections from the hypoglossal motor nuclei of 6 rats and did not see any vacuoles (data not shown). This is neither surprising nor concerning, however, given that the mechanism of cell death is different between SOD1G93A mice and the inducible model we describe here. Patients with ALS typically do not have vacuoles either, and the mechanism of cell death appears to be via apoptosis even in those with SOD1 mutations, in contrast to the prolonged necrotic-like cell death seen in mice that overexpress human SOD1 mutations (Wong et al., 1995; Martin, 2011).

Are the swallowing metrics studied in rodents translational?

Although our CTB-SAP model closely mimics aspects of the ALS mouse model, it is difficult to compare either species directly to human patients. VFSS is considered the gold standard in diagnosing dysphagia, but most studies consist of qualitative descriptions of soft-tissue structures (Martin-Harris and Jones, 2008) that are not readily visible in small rodents (Lever et al., 2015a). Two of the most pervasive aspects of dysphagia in ALS are an impaired ability to manipulate the bolus with the tongue and a delay in triggering the swallowing reflex (Kawaii et al., 2003; Higo et al., 2004; Kühnlein et al., 2008; Jani and Gore, 2016; Hiraoka et al., 2017). Our CTB-SAP treated rats developed a decreased lick rate, which likely corresponds to a decrease in tongue mobility similar to that seen in ALS patients. In addition, the deficit we noted in swallowing rate could be the result of an increased oral transit time as a consequence of impaired tongue function.

It has been suggested that when patients can no longer protrude the tip of their tongue beyond the incisors, diet modifications must be made (Hillel and Miller, 1989). Magnetic resonance images of ALS patients show tongue placement is altered, as demonstrated by the tongue being positioned away from the roof of the mouth and towards the back of the throat even at rest (Cha and Patten, 1989), which appears to occur in our CTB-SAP treated rats (Fig. 5). Our observations are consistent with what has been shown in ALS patients in that maximum tongue protrusion is decreased to the point where the tongue fails to extend beyond the incisors (Fig. 5), and we are currently working on establishing methods to accurately quantify tongue position to optimize translatability of this outcome measure across species.

Lastly, the behavioral changes we observed in our rat model (Fig. 5) are consistent with what is seen in ALS patients with dysphagia, but the reasons for these compensatory behaviors are unclear. For example, we are uncertain why the CTB-SAP treated rats hold their head in a more vertical position while drinking. The position is reminiscent of the “dropped head syndrome” sometimes seen in ALS patients, (Gourie-Devi et al., 2003). In addition, patients with dysphagia are sometimes advised to tuck their chin to reduce the risk of aspiration pneumonia (Ashford et al., 2009; Saconato et al., 2016), but dysphagia does not lead to aspiration in mice (Lever et al., 2015b) and we did not observe aspiration in the CTB-SAP treated rats (data not shown). It is possible this behavior (vertical head position) in the rats is secondary to sensory/proprioceptive effects of our intralingual injections. Previous studies have indicated that intralingual injections of retrograde tracers target neurons in the dorsal root ganglia of the upper cervical spinal cord (C2 and C3) as well as the superior vagal (jugular) and trigeminal ganglia (O’Reilly and Fitzgerald, 1990; Sherif et al., 1991); however, we have not yet examined these ganglia in our model.

Minimal non-targeted cell death after CTB-SAP

We also found CTB(+) neurons in the trigeminal, facial, and superior salivatory motor nuclei after intralingual injection of CTB-SAP; however, only trigeminal motor neuron counts were significantly decreased (Fig. 8). This is consistent with what has been found in the SOD1G93A model, in which significant neurodegeneration of the trigeminal motor nucleus was observed (Robbins et al., 2015). Though systematic pathological investigations of cranial motor nuclei in ALS patients are sparse, consensus among available studies is that the hypoglossal neurons are the most severely affected; trigeminal neurons are also affected, but to a lesser degree (Lawyer and Netsky, 1953; Bonduelle, 1975; Hughes, 1982; DePaul et al., 1988). In previous animal studies, horseradish peroxidase injected into the blade of the tongue or to the transected hypoglossal nerve was retrogradely transported not only to the hypoglossal nucleus but also to the facial motor nucleus and the reticular formation (where dual staining revealed that cells in the reticular formation were a part of the salivatory nucleus; O’Reilly and Fitzgerald, 1990). They did not report staining in the trigeminal motor nucleus, but we suggest this is because horseradish peroxidase was injected into the blade of the tongue itself whereas we targeted the genioglossus with our injections. We consistently saw CTB(+) cells in compartments of the hypoglossal nucleus that supply the intrinsic tongue muscles as well as the geniohyoid, which we attributed to CTB diffusing through the interwoven muscles of the tongue base. In addition, the mylohyoid and anterior digastric muscles lie just ventral to the geniohyoid. We suggest these sublingual muscles, which are innervated by the trigeminal nerve, are also exposed to diffused CTB that results in off-target labeling within the trigeminal motor nucleus. The mylohyoid and anterior digastric muscles assist in jaw opening but primarily function to elevate the hyoid during swallowing. It thus seems likely that loss of innervation to these muscles may also be contributing to dysphagia in these rats. The trigeminal also supplies motor function to the masseter, temporalis, and pterygoid muscles. These masticatory muscles are critical for opening (lateral pterygoid) and closing (masseter, temporalis, and medial pterygoid) the jaw. Lick rate and jaw opening/closing are intertwined and have been shown to correspond 1:1 (Lever et al., 2015a); thus, if the masticatory muscles are weakened by our injections, a deficit in lick rate not related to impaired swallowing function could be observed. We do not believe these masticatory muscles are impacted, however, because the CTB(+) neurons we noted in the trigeminal motor nucleus were all in the ventromedial part of the nucleus (Fig. 8). The ventromedial compartment has been reported to contain neurons that innervate the mylohyoid and anterior digastric muscles, while the dorsolateral compartment contains those for the masseter, temporalis, and pterygoid muscles (Lynch, 1985). In addition, the sublingual muscles are located in greater proximity to the injection site and thus seem more likely to uptake CTB-SAP. Nonetheless, we cannot completely discount the possibility that the masticatory muscles are affected, and are now beginning to study the effects of CTB-SAP on mastication.

Significance

Intralingual injections of CTB-SAP into the genioglossus muscle result in hypoglossal motor deficits (i.e., decreased motor output and motor neuron survival) and dysphagia very similar to what is seen in the SOD1G93A mouse model. However, these deficits occurred without the added complications of other clinical signs such as limb paralysis, as observed in SOD1G93A mice. Perhaps most importantly, the rate and degree of neuronal loss was consistently controlled in our model, which could be advantageous to test new therapies. The pathogenesis of motor neuron death by CTB-SAP is different than in ALS, which does put some limitations on the therapies that can be tested. However, we are primarily interested in studying plasticity in the surviving hypoglossal motor neurons, thus this new model should be ideal for that purpose. For example, SOD1G93A rats show enhanced long-term facilitation (plasticity) in the surviving phrenic motor neurons at disease end-stage (Nichols et al., 2013; 2015a), and we are currently using intrapleural injections of CTB-SAP (Nichols et al., 2015b) to elucidate the mechanisms by which this occurs. It could be similarly hypothesized that intralingual injections of CTB-SAP will result in enhanced plasticity in the surviving hypoglossal motor neurons. Although we did not use markers of apoptosis and instead relied on the appearance of the neurons to gauge viability, it is possible that some neurons exposed to SAP were not yet showing obvious signs and were counted as healthy. Thus, another limitation is that although we see many aspects mimicked here, some of the surviving neurons not showing obvious signs may impact the amount of plasticity. However, we find a similar amount of motor neuron death around the same time frame in our CTB-SAP respiratory model, and these rats exhibit enhanced plasticity (Nichols et al., 2015b; 2017). Thus, we believe this will also likely be the case with hypoglossal plasticity in rats intralingually injected with CTB-SAP. If this hypothesis holds true, the underlying mechanism of enhanced hypoglossal plasticity could be harnessed to preserve swallowing function, which could significantly improve the quality of life of ALS patients with dysphagia.

Finally, the hypoglossal nerve also plays an important role in upper airway control. Hypoglossal motor neurons respond to central and peripheral chemoreceptor activation to move the tongue as needed to dilate and constrict the upper airway (Fuller et al., 1998), and electrical stimulation of the hypoglossal nerve has been shown to be a beneficial treatment for patients with obstructive sleep apnea (Bisogni et al., 2017). Thus, it is possible that this model may also have upper airway effects and may therefore be a relevant platform for researchers working on a variety of conditions associated with hypoglossal motor neuron pathology, including obstructive sleep apnea.

Highlights:

  • 1)

    Intralingual CTB-SAP causes selective death of hypoglossal motor neurons.

  • 2)

    Hypoglossal motor output is reduced in response to intralingual CTB-SAP.

  • 3)

    Intralingual CTB-SAP results in deficits in both lick rate and swallow rate.

  • 4)

    These deficits recapitulate the dysphagia seen in the SOD1G93A mouse model of ALS.

Acknowledgements:

The authors thank Scott Brown and Sarah Stern for assistance with immunohistochemistry and VFSS experiments, Safraaz Mahammed for the custom-designed computer program used for in vivo neurophysiology data analysis, and the Dalton Imaging Core Facilities for the use of the confocal microscope. This work was supported by National Institutes of Health K99/R00 HL119606 (NLN), University of Missouri College of Veterinary Medicine Committee on Research (NLN and TEL), University of Missouri Research Board (NLN and TEL), a Richard Wallace Faculty Incentive grant provided by the Mizzou Alumni Association (NLN), and the Veterinary Research Scholars Program (LAL).

Abbreviations:

ALS

amyotrophic lateral sclerosis

CTB

cholera toxin B

CTB-SAP

cholera toxin B conjugated to saporin

PETCO2

partial pressure of end-tidal carbon dioxide

PaCO2

partial pressure of arterial carbon dioxide

PaO2

partial pressure of arterial oxygen

VFSS

videofluoroscopic swallowing study

Footnotes

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References

  1. Aldes LD (1995) Subcompartmental organization of the ventral (protrusor) compartment in the hypoglossal nucleus of the rat. J Comp Neurol 353(1), 89–108. [DOI] [PubMed] [Google Scholar]
  2. Ashford J, McCabe D, Wheeler-Hegland K, Frymark T, Mullen R, Musson N, Schooling T, Hammond CS (2009) Evidence-based systematic review: Oropharyngeal dysphagia behavioral treatments. Part III--impact of dysphagia treatments on populations with neurological disorders. J Rehabil Res Dev 46(2): 195–204. [PubMed] [Google Scholar]
  3. Bach KB, Mitchell GS (1996) Hypoxia-induced long-term facilitation of respiratory activity is serotonin dependent. Respir Physiol 104(2–3): 251–260. [DOI] [PubMed] [Google Scholar]
  4. Behan M, Moeser AE, Thomas CF, Russell JA, Wang H, Leverson GE, Connor NP (2012) The effect of tongue exercise on serotonergic input to the hypoglossal nucleus in young and old rats. J Speech Lang Hear Res 55(3): 919–929. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bisogni V, Pengo MF, De Vito A, Maiolino G, Rossi GP, Moxham J, Steier J (2017) Electrical stimulation for the treatment of obstructive sleep apnoea: a review of the evidence. Expert Rev Respir Med 11(9): 711–720. [DOI] [PubMed] [Google Scholar]
  6. Bonduelle M (1975) Amyotrophic lateral sclerosis In: Handbook of clinical neurology, vol. 22 (Vincken PJ, Bruyn GW, DeJong JM, eds), pp 281–338. New York: Elsevier. [Google Scholar]
  7. Cha CH, Patten BM (1989) Amyotrophic lateral sclerosis: Abnormalities of the tongue on magnetic resonance imaging. Ann Neurol 25: 468–472. [DOI] [PubMed] [Google Scholar]
  8. Corcia P, Pradat P-F, Salachas F, Bruneteau G, Le Forestier N, Seilhean D, Hauw JJ, Meininger V (2008) Causes of death in a post-mortem series of ALS patients. Amyotroph Lateral Scler 9: 59–62. [DOI] [PubMed] [Google Scholar]
  9. DePaul R, Abbs JH, Caligiuri M, Gracco VL, Brooks BR (1988) Hypoglossal, trigeminal, and facial motoneuron involvement in amyotrophic lateral sclerosis. Neurology 38: 281–283. [DOI] [PubMed] [Google Scholar]
  10. ElMallah MK, Falk DJ, Lane MA, Conlon TJ, Lee KZ, Shafi NI, Reier PJ, Byrne BJ, Fuller DD (2012) Retrograde gene delivery to hypoglossal motoneurons using adenoassociated virus serotype 9. Hum Gene Ther Methods 23(2): 145–156. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Fuller D, Mateika JH, Fregosi RF (1998). Co-activation of tongue protrudor and retractor muscles during chemoreceptor stimulation in the rat. J Physiol 507 (Pt. 1): 265–276. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Gourie-Devi A, Nalini A, Sandhya S (2003) Early or late appearance of "dropped head syndrome" in amyotrophic lateral sclerosis. J Neurol Neurosurg Psychiatry 74(5): 683–686. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Gurney ME, Pu H, Chiu AY, Dal Canto MC, Polchow CY, Alexander DD, Caliendo J, Hentati A, et al. , (1994) Motor neuron degeneration in mice that express a human Cu,Zn superoxide dismutase mutation. Science 264(5166): 1772–1775. [DOI] [PubMed] [Google Scholar]
  14. Hadjikoutis S, Wiles CM (2001) Respiratory complications related to bulbar dysfunction in motor neuron disease. Acta Neurol Scand 103: 207–213. [PubMed] [Google Scholar]
  15. Higo R, Tayama N, Nito T (2004) Longitudinal analysis of progression of dysphagia in amyotrophic lateral sclerosis. Auris Nasus Larynx 31(3):247–54. [DOI] [PubMed] [Google Scholar]
  16. Hillel AD, Miller R (1989) Bulbar amyotrophic lateral sclerosis: patterns of progression and clinical management. Head Neck 11(1): 51–59. [DOI] [PubMed] [Google Scholar]
  17. Hiraoka A, Yoshikawa M, Nakamori M, Hosomi N, Nagasaki T, Mori T, Oda M, Maruyama H, et al. , (2017) Maximum tongue pressure is associated with swallowing dysfunction in ALS patients. Dysphagia 32(4): 542–547. [DOI] [PubMed] [Google Scholar]
  18. Hoffman MS, Nichols NL, Macfarlane PM, Mitchell GS (2012) Phrenic long-term facilitation following acute intermittent hypoxia requires spinal ERK activation but not TrkB synthesis. J Appl Physiol 113: 1184–1193. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Hughes JT (1982) Pathology of amyotrophic lateral sclerosis In: Human motor neuron diseases (Rowland LP, ed), pp 61–74. New York: Raven Press. [Google Scholar]
  20. Jani MP, Gore GB (2016) Swallowing characteristics in amyotrophic lateral sclerosis. NeuroRehabilitation 39: 273–276. [DOI] [PubMed] [Google Scholar]
  21. Kawai S, Tsukuda M, Mochimatsu I, Enomoto H, Kagesato Y, Hirose H, Kuroiwa Y, Suzuki Y (2003) A study of the early stage of dysphagia in amyotrophic lateral sclerosis. Dysphagia 18(1): 1–8. [DOI] [PubMed] [Google Scholar]
  22. Kiernan MC, Vucic S, Cheah BC, Turner MR, Eisen A, Hardiman O, Burrell JR, Zoing MC (2011) Amyotrophic lateral sclerosis Lancet 377: 942–955. [DOI] [PubMed] [Google Scholar]
  23. Kong J, Xu Z (1998) Massive mitochondrial degeneration in motor neurons triggers the onset of amyotrophic lateral sclerosis in mice expressing a mutant SOD1. J Neurosci 18(9): 3241–3250. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Kühnlein P, Gdynia HJ, Sperfeld AD, Lindner-Pfleghar B, Ludolph AC, Prosiegel M, Riecker A (2008) Diagnosis and treatment of bulbar symptoms in amyotrophic lateral sclerosis. Nat Rev Neurol 4(7): 366. [DOI] [PubMed] [Google Scholar]
  25. Kurian KM, Forbes RB, Colville S, Swingler RJ (2009) Causes of death and clinical grading criteria in a cohort of amyotrophic lateral sclerosis cases undergoing autopsy from the Scottish Motor Neurone Disease Register. J Neurol Neurosurg Psychiatry 80: 84–87. [DOI] [PubMed] [Google Scholar]
  26. Langmore SE, Lehman ME (1994) Physiologic deficits in the orofacial system underlying dysarthria in amyotrophic lateral sclerosis. J Speech Hear Res 37: 28–37. [DOI] [PubMed] [Google Scholar]
  27. Lawyer T, Netsky MG (1953) Amyotrophic lateral sclerosis: a clinicoanatomic study of fifty-three cases. Arch Neurol Psychiatr 69: 171–192. [DOI] [PubMed] [Google Scholar]
  28. Lee KZ, Qiu K, Sandhu MS, Elmallah MK, Falk DJ, Lane MA, Reier PJ, Byrne BJ, Fuller DD (2011) Hypoglossal neuropathology and respiratory activity in pompe mice. Front Physiol 2: 31. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Lencer WI, Tsai B (2003) The intracellular voyage of cholera toxin: going retro. Trends in biochemical sciences 28(12): 639–45. [DOI] [PubMed] [Google Scholar]
  30. Lever TE, Braun SM, Brooks RT, Harris RA, Littrell LL, Neff RM, Hinke CJ, Allen MJ, Ulsas MA (2015a) Adapting human videofluoroscopic swallow study methods to detect and characterize dysphagia in murine disease models. J Vis Exp 97: e52319. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Lever TE, Brooks RT, Thombs LA, Littrell LL, Harris RA, Allen MJ, Kadosh MD, Robbins KL (2015b) Videofluoroscopic validation of a translational murine model of presbyphagia. Dysphagia 30(3): 328–342. [DOI] [PubMed] [Google Scholar]
  32. Lever TE, Gorsek A, Cox KT, O’Brien KF, Capra NF, Hough MS, Murashov AK (2009) An animal model of oral dysphagia in amyotrophic lateral sclerosis. Dysphagia 24(2): 180–195. [DOI] [PubMed] [Google Scholar]
  33. Lever TE, Simon E, Cox KT, Capra NF, O’Brien KF, Hough MS, Murashov AK (2010) A mouse model of pharyngeal dysphagia in amyotrophic lateral sclerosis. Dysphagia 25(2): 112–126. [DOI] [PubMed] [Google Scholar]
  34. Lian T, Ho RJ (1997) Cholera toxin B-mediated targeting of lipid vesicles containing ganglioside GM1 to mucosal epithelial cells. Pharm Res 14(10): 1309–1315. [DOI] [PubMed] [Google Scholar]
  35. Llewellyn-Smith IJ, Martin CL, Arnolda LF, Minson JB (1999) Retrogradely transported CTB-saporin kills sympathetic preganglionic neurons. Neuroreport 10(2): 307–12. [DOI] [PubMed] [Google Scholar]
  36. Llewellyn-Smith IJ, Martin CL, Arnolda LF, Minson JB (2000) Tracer toxins: cholera toxin B-saporin as a model. J Neurosci Methods 103(1): 83–90. [DOI] [PubMed] [Google Scholar]
  37. Lujan HL, Palani G, Peduzzi JD, DiCarlo SE (2010) Targeted ablation of mesenteric projecting sympathetic neurons reduces the hemodynamic response to pain in conscious, spinal cord-transected rats. Am J Physiol Regul Integr Comp Physiol 298(5): 1358–1365. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Lynch R (1985) A qualitative investigation of the topographical representation of masticatory muscles within the motor trigeminal nucleus of the rat: a horseradish peroxidase study. Brain research 327(1–2): 354–8. [DOI] [PubMed] [Google Scholar]
  39. Martin LJ (2011) Mitochondrial pathobiology in ALS. J. Bioenerg. Biomembr. 43(6): 569–579. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Martin-Harris B, Jones B (2008) The videofluorographic swallowing study. Phys Med Rehabil Clin N Am 19: 769–785. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. McClung JR, Goldberg SJ (1999) Organization of motoneurons in the dorsal hypoglossal nucleus that innervate the retrusor muscles of the tongue in the rat. Anat Rec 254(2), 222–230. [DOI] [PubMed] [Google Scholar]
  42. Nichols NL, Craig TA, Tanner MA (2017) Phrenic long-term facilitation following intrapleural CTB-SAP-induced respiratory motor neuron death. Respir physiol neurobiol. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Nichols NL, Dale EA, Mitchell GS (2012) Severe acute intermittent hypoxia elicits phrenic long-term facilitation by a novel adenosine-dependent mechanism. J Appl Physiol 112: 1678–1688. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Nichols NL, Gowing G, Satriotomo I, Nashold LJ, Dale EA, Suzuki M, Avalos P, Mulcrone PL, et al. , (2013) Intermittent hypoxia and stem cell implants preserve breathing capacity in a rodent model of amyotrophic lateral sclerosis. Am J Respir Crit Care Med 187(5): 535–542. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Nichols NL, Satriotomo I, Harrigan DJ, Mitchell GS (2015a) Acute intermittent hypoxia induced phrenic long-term facilitation despite increased SOD1 expression in a rat model of ALS. Exper Neurol 273: 138–150. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Nichols NL, Vinit S, Bauernschmidt L, Mitchell GS (2015b) Respiratory function after selective respiratory motor neuron death from intrapleural CTB-saporin injections. Exper Neurol 267: 18–29. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Onesti E, Schettino I, Gori MC, Frasca V, Ceccanti M, Cambieri C, Ruoppolo G, Inghilleri M (2017) Dysphagia in amyotrophic lateral sclerosis: Impact on patient behavior, diet adaptation, and riluzole management. Front Neurol 8: 94. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. O’Reilly PMR, Fitzgerald MJT (1990) Fibre composition of the hypoglossal nerve in the rat. J Anat 172: 227–243. [PMC free article] [PubMed] [Google Scholar]
  49. Paxinos G, Watson C (2005) The Rat Brain in Stereotaxic Coordinates, 5th ed. Elsevier. [DOI] [PubMed] [Google Scholar]
  50. Perrin C, Unterborn JN, d’Ambrosio C, Hill NS (2004) Pulmonary complications of chronic neuromuscular diseases and their management. Muscle Nerve 29: 5–27. [DOI] [PubMed] [Google Scholar]
  51. Robbins K, Allen M, Lever T (2015) Low copy number SOD1-G93A mice are better suited for dysphagia research compared to the high copy number model. Amyotrophic Lateral Sclerosis and Frontotemporal Degeneration 16: 217–226.26517034 [Google Scholar]
  52. Saconato M, Chiari BM, Lederman HM, Gonçalves MIR (2016) Effectiveness of Chin-tuck Maneuver to Facilitate Swallowing in Neurologic Dysphagia Int Arch Otorhinolaryngol 20(1): 13–17. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Sherif MF, al-Zuhair AG, Kharbat BA (1991) Identification of sensory neurons supplying receptors in lingual muscles of the rat: histochemical and retrograde labeling study with horseradish peroxidase. Histol Histopathol 6(4): 549–558. [PubMed] [Google Scholar]
  54. Tomik B, Guiloff RJ (2010) Dysarthria in amyotrophic lateral sclerosis: A review. Amyotroph Lateral Scler 11: 4–15. [DOI] [PubMed] [Google Scholar]
  55. Wong PC, Pardo CA, Borchelt BR, Lee MK, Copeland NG, Jenkins NA, Sisodia SS, Cleveland DW, Price DL (1995). An adverse property of a familial ALS-liked SOD1 mutation causes motor neuron disease characterized by vacuolar degeneration of mitochondria. Neuron 14(6): 1105–1116. [DOI] [PubMed] [Google Scholar]

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