This work was funded by a Spinal Cord Injury/Disease Research Program Grant from the Missouri Spinal Cord Injury/Disease Research Program (NLN and WDA).

All procedures were approved and conducted in compliance with the guidelines established by the Institutional Animal Care and Use Committee at the University of Missouri. Experiments were performed on adult male Sprague-Dawley rats, aged 11 to 15 weeks. These rats were housed in pairs and kept under a 12:12 light-dark cycle, with access to standard commercial pelleted food and HCl-treated water available at all times.
1. Animal preparation and anesthesia delivery
<ol>
	<li>Wear suitable personal protection equipment while handling rats.</li>
	<li>Administer inhalation anesthetic with 3-5% isoflurane, ensuring proper induction. Once the rat is adequately anesthetized, place it in a supine position and maintain the anesthesia with 1-3% inhaled isoflurane. Check the sufficiency of the anesthesia depth by gently applying pressure to the hindlimb footpad using forceps to ensure there is no withdrawal response. 
	NOTE: Based on the rat&#39;s size and weight, monitor the depth of anesthesia, and adjust the isoflurane concentration accordingly.</li>
	<li>Maintain the temperature of 37 &#176;C using a thermostatic warming plate to prevent variations in temperature that can impact CMAP amplitude and latency.</li>
	<li>Apply a veterinary petroleum-based ointment to the eyes to prevent dryness. Monitor the depth of anesthesia by observing the respiration rate and checking for withdrawal responses upon applying pressure to the footpad with forceps.</li>
	<li>Remove hair from the lower third of the chest and neck to be studied using clippers. Monitor the respiration of the rat during the entire experiment. 
	NOTE: Following the CMAP and MUNE recordings and discontinuation of anesthesia, do not leave the rat unattended until it has regained sufficient consciousness. Do not return the animal to the home cage until fully recovered.</li>
</ol>
2. Electrode placement and setup
<ol>
	<li>Place a pair of 28 G monopolar needle electrodes to record the CMAP, SMUP, and MUNE as depicted in Figure 1.
	<ol>
		<li>Place the active (E1) needle electrode subcutaneously over the mid clavicular line inferior to the last rib border, and the reference (E2) needle electrode subcutaneously in the angle between the xyphoid process and last sternocostal cartilage. 
		NOTE: The needle electrodes should not be inserted into the diaphragm muscle; instead, they should be positioned in the subcutaneous area.</li>
	</ol>
	</li>
	<li>For stimulation of the phrenic nerve at the carotid sheet, use a pair of 28 G monopolar needle electrodes as the cathode and anode for nerve stimulation and placed subcutaneously over the lateral neck between the anterior and middle scalene muscles, separated by approximately 1 cm. Ensure that the placement of the stimulating needles is at the level below the fourth cervical vertebrae (C4). 
	NOTE: Avoid inserting the stimulating electrodes too deep to avoid injury of the phrenic nerve or other structure. Figure 1 illustrates electrode placement.</li>
	<li>For the ground electrode, place a disposable surface electrode on the tail.</li>
</ol>
3. Data acquisition
<ol>
	<li>Phrenic CMAP
	<ol>
		<li>Record phrenic CMAP responses by applying monophasic cathodic square-wave pulses with a duration of 0.1 ms and intensity ranging from 60 to 100 mA to stimulate the phrenic nerve.</li>
		<li>Obtain CMAP responses while progressively increasing the stimulus intensity until the amplitude of the response ceases to show any further increase. To ensure supramaximal stimulation, raise the stimulus intensity to approximately 120% of the level used to elicit a maximal response, and record an additional response. If the CMAP size no longer increases, consider this response as the maximum CMAP. 
		NOTE: Delivering stimulation during exhalation is preferable to minimize concurrent muscle activity noise during CMAP recording.</li>
		<li>Measure and document the peak-to-peak amplitudes of the CMAP in millivolts (mV) (Figure 2). 
		NOTE: CMAP amplitude can be assessed base-to-peak and peak-to-peak. Clinical electrodiagnostic systems are often defaulted to assess base-to-peak which is calculated from the isoelectric baseline to the initial negative peak.</li>
	</ol>
	</li>
	<li>Average single motor unit potential (SMUP) size and MUNE calculation
	<ol>
		<li>Calculate the average SMUP size using an incremental stimulation technique.
		<ol>
			<li>To elicit incremental responses, administer submaximal stimulation with a duration of 0.1 ms at 1 Hz frequency, gradually increasing the intensity in 0.03 mA increments until a minimal all-or-none response is achieved. Acquire the initial response with a stimulus intensity ranging between 2 mA and 10 mA.</li>
			<li>If the initial response does not occur with a stimulus intensity between 2 mA and 10 mA, modify the position of the stimulating cathode, either bringing it closer or moving it farther from the phrenic nerve in the neck, to decrease or increase the necessary stimulus intensity, respectively.</li>
			<li>If the first incremental response is achieved with a stimulus intensity ranging from 2 mA to 10 mA, save the first response and acquire additional increments with progressively higher stimulus intensities, adjusting in increments of 0.03 mA, to achieve a total of 9 additional increments that fulfill the following criteria in step 3.2.2. 
			NOTE: Each SMUP is quantified by subtracting each increment from the preceding increment.</li>
		</ol>
		</li>
		<li>While measuring the incremental responses, make sure that each increment meets the following criteria:
		<ol>
			<li>Make sure that the initial negative peak of the incremental responses aligns temporally with the negative peak of the maximal CMAP response. 
			NOTE: The slight movements observed due to background noise from respiratory cycles are inherent to the nature of the experiment. However, the consistent presence of SMUPs during live observation confirms their identity for that specific CMAP.</li>
			<li>As the diaphragm is a dynamic muscle involved in respiration, each breathing cycle may induce baseline movement. Thus, verify the stability and absence of fractionation in each incremental response by confirming consistency across three duplicate responses. 
			NOTE: To distinguish low-amplitude evoked potentials, especially the first SMUP, from background noise due to respiratory activity, it is important to remain vigilant and observe for 3-4 respiratory cycles. Also ensure the alignment of peaks with those of the CMAP for accuracy.</li>
			<li>Ensure that each increment is distinct and larger than the previous one. Therefore, visually distinguish incremental responses in real time, observing them as they overlay on the previously recorded increments. 
			NOTE: Multiple replicates of the stimulus at each amplitude might be conducted to ensure consistency in the increments and compliance with the predefined criteria.</li>
			<li>After visually verifying each increment with the aforementioned criteria, confirm that the increment amplitude is at least 25 &#181;V. If the increment is below 25 &#181;V, discard the measurement and re-assess the response.</li>
			<li>Following the recording of 10 incremental responses, verify that the amplitude of each increment response is not greater than one-third of the combined amplitude of all 10 increments, representing the total amplitude of the final response. If this criterion is not satisfied, repeat the measurement of the 10 incremental responses. 
			NOTE: The threshold of one-third is based on the assumption that each incremental response represents the activation of a single motor unit. If the amplitude of any incremental response exceeds one-third of the combined amplitude of all ten increments, it suggests that the response may not be solely attributable to the activation of a single motor unit. Instead, it could be influenced by the recruitment of additional motor units or the presence of non-specific activity, such as electrical noise or artifacts22,26.</li>
			<li>To estimate the average amplitude of the SMUPs, average the values of the 10 increments. Another method to measure average SMUP amplitude is to divide the entire amplitude of the final incremental response by the total number of increments11.</li>
		</ol>
		</li>
		<li>Determine MUNE by dividing the maximum CMAP amplitude (peak-to-peak) by the average SMUP amplitude (peak-to-peak). In certain electrophysiological systems, SMUPs are recorded in microvolts (&#181;V), while CMAP is usually expressed in millivolts (mV). If required, convert CMAP and SMUP measurements to the same units before calculating MUNE. 
		NOTE: Peak-to-peak CMAP, average SMUP, and MUNE are typically automatically calculated by clinic electromyography systems): 
		<img alt="Equation 1" src="/files/ftp_upload/66568/66568eq01v2.jpg" style="margin: auto;" /> 
		CMAP = Compound Muscle Action Potential 
		SMUP = Single Motor Unit Potential 
		MUNE = Motor Unit Number Estimation</li>
	</ol>
	</li>
</ol>

Quantifying Phrenic Motor Unit Function in Rodent Models of Neurodegeneration

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WDA has received research funding from NMD Pharma, Avidity Biosciences, and consulting fees from NMD Pharma, Avidity Biosciences, Dyne Therapeutics, Novartis, Design Therapeutics, Catalyst Pharmaceuticals, and Novartis.

In MN degenerative diseases, such as ALS, it is crucial to assess the MUs involved in ventilation28. Despite the occurrence of respiratory MN degeneration in ALS patients, the specific onset and progression of MN death remain incompletely understood29,30,31. Recognizing the significance of this aspect, various models, both genetic-based (e.g., SOD12,32) and non-genetic-based (e.g., CTB-SAP intrapleural injection3,27,33), have been employed to emulate respiratory impairment in animal models of neurodegenerative diseases. Accordingly, identifying a biomarker for diagnosing, monitoring, and assessing potential treatment effects is advantageous for these models. Additionally, biomarkers can facilitate the clinical translation of preclinical research findings.
To quantify the number of innervating MNs in rodent degenerative models, different labeling techniques, including retrograde tracers and adenoviruses, have been employed25,34,35,36. Previously, immunohistochemistry has been utilized to label phrenic motor neurons in the anterior horn of the cervical spinal cord2,3,33. Labeling techniques, though valuable for MN evaluations, have limitations in assessing the functionality of MUs and are not suitable for longitudinal assessments37. Objective electrophysiological assessments are crucial for successful cross-species translation, especially considering pronounced structural differences in the functional and anatomical features of the motor systems between humans and rodents38,39. Incorporating neurobiological differences presents a challenge in translating findings from rodent models to patients. MUNE overcomes these obstacles by serving as an objective electrophysiological measure of MN function, allowing the evaluation of functional MU connectivity as a biomarker in MN degenerative model experiments3,40.
CMAP, SMUP, and MUNE are commonly employed in research studies and for evaluating patients with neuromuscular disorders9,10,11. These potential biomarkers are minimally invasive, enabling longitudinal assessment of function within the same individual over time. Importantly, while they do not directly gauge the activation or recruitment of the MU by cortical MNs, they do provide a clinically relevant estimation of the MN integrity and its functional counterpart, the MU. Rodent models of MN degenerative diseases are crucial for comprehending the physiopathologic mechanisms underlying human diseases and for the preclinical advance of effective treatments. Developing outcome measures and biomarkers that are translatable across species can streamline and expedite the translation of promising preclinical findings to clinical trials11. Given the relative complexity of the measures, we have adapted and refined these techniques from rat limbs to the diaphragm muscle. This translation has been presented in a visual format to facilitate broader utilization and implementation in rat studies.
From our experience, the clinical electrodiagnostic systems are well-suited for the studies outlined in this context. This suitability stems from enhanced ergonomics in the interface between the examiner and the electrodiagnostic system, facilitating convenient control. In our laboratory group, we utilize a two-channel system featuring two non-switched amplifier channels. These channels are equipped with a 24-bit Analog to Digital Converter and operate at a sampling rate of 48 kHz per channel. The hardware gain is adjustable within a range from 10nV to 100 mV per division. We employed a low-frequency filter with a range spanning from 0.2 Hz to 5 kHz, and the high-frequency filter settings covered a range from 30 Hz to 10 kHz. Additionally, we utilized a constant-current stimulator with adjustable intensity ranging from 0 to 100 mA and duration from 0.02 to 1 ms. The majority of clinical systems offer comparable features that are suitable for recording CMAP, SMUP, and MUNE responses. These systems can be adjusted accordingly to ensure accurate recording of the desired data. For instance, CMAP amplitude can be assessed base-to-peak and peak-to-peak. Clinical electrodiagnostic systems are often defaulted to assess base-to-peak which is calculated from the isoelectric baseline to the initial negative peak.
The process of obtaining CMAP, SMUP, and MUNE responses from the diaphragm involves several crucial steps. Consistent and accurate electrode placement, along with adequate electrode depth, is essential for reliably measuring amplitudes and minimizing background noises during recording. Incorrect placement of stimulating electrodes can lead to unintended consequences. Placing stimulating electrodes too posteriorly can result in exaggerated forelimb muscle contractions to brachial plexus excitation, while higher than C4 level placement can make stimulation of the phrenic nerve more difficult. Deeper placement carries the risk of injuring the carotid artery or evoking the vagus nerve, potentially causing cardiac conductivity issues. Placing recording electrodes higher or lower than the last rib, measurements from intercostal or rectus abdominis stimulation may occur, respectively. To avoid recording from other muscles, notice that the diaphragm negative peak of CMAP latency is typically 1.5-3.5 ms, and its characteristic shape is shown in Figure 2. Our observations indicated that needle electrodes yield more consistent CMAP, SMUP, and MUNE recordings compared to disc-shaped electrodes for diaphragm muscle. This enhanced consistency is attributed to the dome shape of the diaphragm muscle and the presence of other muscles in proximity, such as the intercostal and rectus abdominis muscles. Additionally, flexible placement of the needle electrodes is another advantageous factor. Regarding electrical stimulation intensity, we utilized a range of 60 to 100 mA to stimulate the phrenic nerve, which exceeds the reported ranges for other motor systems. This difference arises from the deeper location of the phrenic nerve in the neck compared to the more superficial locations of the sciatic nerve and brachial plexus in the hindlimb and forelimb, respectively.
Obtaining the average SMUP poses greater technical challenges compared to CMAP. The smaller response size, in the microvolt range rather than millivolts, makes the impact of background noise more pronounced. To mitigate background noise, consider optimizing electrode placement by adjusting the ground electrode, cathode, and anode. Additionally, ensure there is minimal interference from nearby electrical devices in the experimental setup. A Faraday cage, commonly utilized in intracellular electrophysiology applications, is not necessary for this setup. The visual identification of individual SMUP responses is a challenging skill that requires practice for consistent and repeatable results. As the diaphragm is a dynamic muscle, the baseline can shift with each ventilation cycle. Being attentive to the SMUP increments ensuring alignment with the previous SMUP is a critical factor in overcoming this challenge. Additionally, ensuring that the SMUPs initiate within the latency timeframe of the maximal CMAP is important for accurate recordings.
Another challenge of diaphragm recording is the asymmetric nature of this muscle, characterized by varying thicknesses from left to right and influenced by the pressure exerted by the liver on the right side41. To address this, we conducted electrophysiological measurements separately on the left and right hemidiaphragms. Following that, we calculated the averages for CMAP, SMUP, and MUNE responses separately.
Since the CMAP response reflects the collective depolarization of muscle fibers in a specific muscle, any pathology affecting the phrenic MN to the diaphragm muscle fiber can result in a reduction in CMAP size. Hence, calculating the average CMAP from bilateral hemidiaphragms offers an excellent assessment of the overall functional status of the respiratory neuromuscular system. Compensatory changes, such as collateral sprouting, can lead to the preservation of CMAP size even in the presence of MN or motor axonal loss11. Recording individual increments allows for the estimation of the average output of single MUs (SMUP size), presenting more detailed insights into the functional status of MUs of the respiratory system. Hence, the MUNE technique becomes essential for evaluating the phrenic MNs or axons&#39; input to the diaphragm. Applying these biomarkers in preclinical experiments of therapies for studying MN diseases holds the potential to improve the translation to clinical trials.

Phrenic motor neurons (MNs), extending from C3 to C6 myotome levels, form the final link from the central nervous system (CNS) to the diaphragm muscle1. Phrenic motor units (MUs) are comprised of a single spinal MN and its innervated diaphragm muscle fibers forming the smallest functional unit of the respiratory neuromuscular system. The ventilatory function requires adequate contraction of the diaphragm muscle achieved through coordinated activation of the phrenic MU pool2,3. Many neurological diseases, including amyotrophic lateral sclerosis (ALS), result in severe ventilatory impairment, ultimately contributing to the cause of death4.
Several electrophysiological approaches can be employed to evaluate and monitor the integrity of the motor unit (MU) pool in vivo. Compound muscle action potential (CMAP) reflects the summated depolarization of all muscle fibers in a specific muscle or muscle group after peripheral nerve stimulation and is sensitive to a range of neuromuscular conditions, including ALS5,6 and spinal muscular atrophy (SMA)7,8,9. A limitation of CMAP assessment is that collateral sprouting can lead to maintained CMAP amplitude and area even in the presence of MU loss10. To overcome this limitation, modifications have been made to the CMAP technique to evaluate both motor unit number and size11. Additionally, an in vivo study investigating the functional assessment of diaphragm CMAP by an electrophysiological system suggested that it may also be feasible to utilize the described diaphragm CMAP recording technique for motor unit number estimation12.
The incremental motor unit number estimation (MUNE) technique was initially introduced in the early 1970s by McComas et al. for the extensor digitorum brevis muscle in humans13. The incremental MUNE approach was a modification of the traditional CMAP recording technique during which a gradually increasing stimulation was delivered to record quantal, all-or-none submaximal increments as indices of single motor unit responses. The summed and averaged increments were used to calculate an estimate for the size of a single motor unit potential (SMUP). This calculated size was then divided into the CMAP amplitude to estimate the number of MUs innervating the muscle under examination11. MUNE demonstrates high sensitivity in detecting and monitoring motor unit loss, allowing for the identification of motor unit dysfunction before observable changes in measures such as CMAP amplitude or area14,15. In ALS patients, MUNE has proven to be exceptionally sensitive, serving as a prominent biomarker for disease onset, progression, and prognosis16,17.
Numerous adaptations of MUNE have been developed and widely used to assess MU function in conditions such as neurodegeneration, neural injury, and the natural aging process18,19,20,21. Since the initial description, various adaptations utilizing both electrophysiological responses and incremental force (mechanical) measurements have been employed in both human studies and animal models22. MUNE provides a non-invasive functional assessment of motor neuron connectivity with the muscle. Longitudinally applying MUNE enables the understanding of disease or induced phenotype progression and the evaluation of protective or regenerative effects of therapeutic interventions, both in clinical and preclinical settings. Regardless of the effectiveness of MUNE measures reproducibility and the clinical relevance of the technique for MU pools throughout most of the human body, efforts have largely focused on limb muscles in rodent muscles10,23,24,25.
Therefore, the objectives of this study were to describe an approach to obtaining compound muscle action potential (CMAP), SMUP, and phrenic motor unit number (MUNE) as in vivo assessments that can be used longitudinally in preclinical rodent studies to quantify the MUNE, motor unit size (represented as SMUP), and CMAP. Furthermore, we present representative data that highlights the loss of diaphragm MU number following intrapleural administration of a phrenic MN degenerative agent, cholera toxin B fragment conjugated to saporin (CTB-SAP).

<table><tbody><tr><td>2 mL Glass Syringe</td><td>Kent Scientific Corporation</td><td>SOMNO-2ML</td><td></td></tr><tr><td>50 mL, Model 705 RN syringe</td><td>Hamilton Company</td><td>7637-01</td><td>Utilized to conduct intrapleural injection</td></tr><tr><td>5008 - Formulab Diet</td><td>LabDiet</td><td>0001325</td><td></td></tr><tr><td>Autoclavable 26 G needles (26S RN 9.52 mm 40&amp;deg;)</td><td>Hamilton Company</td><td>7804-04</td><td>Utilized to conduct intrapleural injection</td></tr><tr><td>Cholera toxin B-subunit (CTB)</td><td>MilliporeSigma</td><td>C9903</td><td>Utilized for intrapleural injection to label surviving motor neurons</td></tr><tr><td>Cholera toxin B-subunit conjugated to saporin (CTB-SAP)</td><td>Advanced Targeting Systems</td><td>IT-14</td><td>Utilized for intrapleural injection to cause motor neuron death</td></tr><tr><td>Detachable Cable</td><td>Technomed</td><td>202845-0000</td><td>to connect the recorder electrode to the electrodiagnostic machine</td></tr><tr><td>Disposable 2&quot; x 2&quot; disc electrode with leads</td><td>Cadwell</td><td>302290-000</td><td>ground electrode</td></tr><tr><td>disposable monopolar needles 28 G</td><td>Technomed</td><td>202270-000</td><td>cathode and anode stimulating electrodes- recording electrodes</td></tr><tr><td>EMG needle cable (Amp/stim switch box)</td><td>Cadwell</td><td>190266-200</td><td>to connect monopolar electrodes to electrodiagnostic stimulator</td></tr><tr><td>Helping Hands alligator clip with iron base</td><td>Radio Shack</td><td>64-079</td><td>Maintaining recording electrode placement&amp;nbsp;</td></tr><tr><td>Isoflurane (250 mL bottle)</td><td>Piramal Healthcare</td><td></td><td></td></tr><tr><td>monoject curved tip irrigating syringe</td><td>Covidien</td><td>81412012</td><td>utilized for application of electrode gel</td></tr><tr><td>PhysioSuite Physiological Monitoring System with RightTemp Homeothermic Warming</td><td>Kent Scientific Corporation</td><td>PS-RT</td><td>Includes infrared warming pad, rectal probe, and pad temperature probe</td></tr><tr><td>Pro trimmer Pet Grooming Kit</td><td>Oster</td><td>078577-010-003</td><td>clippers for hair removal</td></tr><tr><td>Saporin (SAP)</td><td>Advanced Targeting Systems</td><td>PR-01</td><td>Utilized for intrapleural injection (control agent when injected by itself)</td></tr><tr><td>Sierra Summit EMG system</td><td>Cadwell Industries, Inc., Kennewick, WA</td><td></td><td>portable electrodiagnostic system</td></tr><tr><td>SomnoSuite Low-Flow Digital Anesthesia System</td><td>Kent Scientific Corporation</td><td>SOMNO</td><td>Includes anti-spill, anti-vapor bottle top adapter; Y adapter tubing; charcoal scavenging filter</td></tr><tr><td>Sprague-Dawley rat</td><td>Envigo colony 208a, Indianapolis, IN</td><td></td><td></td></tr><tr><td>Veterinarian petroleum-based ophthalmic ointment&amp;nbsp;</td><td>Puralube</td><td>26870</td><td>applied during anesthesia to avoid corneal injury</td></tr></tbody></table>

assessing rat diaphragm motor unit connectivity outcome measures as quantitative biomarkers of phrenic motor neuron degeneration and compensation

Loss of ventilatory muscle function is a consequence of motor neuron injury and neurodegeneration (e.g., cervical spinal cord injury and amyotrophic lateral sclerosis, respectively). Phrenic motor neurons are the final link between the central nervous system and muscle, and their respective motor units (groups of muscle fibers innervated by a single motor neuron) represent the smallest functional unit of the neuromuscular ventilatory system. Compound muscle action potential (CMAP), single motor unit potential (SMUP), and motor unit number estimation (MUNE) are established electrophysiological approaches that enable the longitudinal assessment of motor unit integrity in animal models over time but have mostly been applied to limb muscles. Therefore, the objectives of this study are to describe an approach in preclinical rodent studies that can be used longitudinally to quantify the phrenic MUNE, motor unit size (represented as SMUP), and CMAP, and then to demonstrate the utility of these approaches in a motor neuron loss model. Sensitive, objective, and translationally relevant biomarkers for neuronal injury, degeneration, and regeneration in motor neuron injury and diseases can significantly aid and accelerate experimental research discoveries to clinical testing.

The CMAP, SMUP, and MUNE techniques outlined in this report enable the recording of neuromuscular function in the diaphragm muscle employing minimally invasive electrode placement (Figure 1). The parameters of amplitude and area can be employed to characterize the supramaximal CMAP size, providing an overall measure of muscle group output (Figure 2). However, in our current methods, we rely on amplitude to quantify both CMAP and SMUP sizes. CMAP, SMUP, and MUNE can be utilized to measure neuromuscular function in various rat models of neuromuscular disease. To demonstrate MU loss in the context of MN degeneration, rats underwent bilateral intrapleural injection with CTB-SAP (25 &#181;g) to target phrenic MNs and extra CTB (25 &#181;g), and control rats received unconjugated CTB (25 &#181;g) and SAP (25 &#181;g)1,27.
In Figure 3, findings in an adult control rat and an adult rat (12 weeks) seven days following intrapleural CTB-SAP injection are compared. Following intrapleural injection of CTB-SAP, MUNE is reduced at 60 estimated functional motor units compared with normal findings of 74 functional motor units in the control rat. However, the CMAP amplitude in the CTB-SAP rat (14.25 mV peak-to-peak) exhibited minimal changes compared to the control (14. 5 mV peak-to-peak) likely due to collateral sprouting. In Figure 4, diaphragm CMAP, SMUP, and MUNE were acquired bilaterally and averaged at baseline/preinjection (n = 14), as well as 7 (n = 14), 14 (n = 14), 21 (n = 6), and 28 (n = 14) days post injection. In Figure 4, no baseline differences between groups were found. MUNE showed a significant change for time (p &#60; 0.05), CTB-SAP (p &#60; 0.001), and interaction time x CTB-SAP (p &#60; 0.01) with ~40% reduction of MUNE in CTB-SAP rats. The average SMUP of CTB-SAP rats demonstrated a significant change with time (p &#60; 0.05) and CTB-SAP (p &#60; 0.01), but no significant interaction with ~50-60% increase in SMUP amplitude. CMAP showed no significant change.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66568/66568fig01.jpg" /> 
Figure 1: Electrode placement. The (A) stimulating cathode and (B) anode are subcutaneously inserted in the lateral neck between the anterior and middle scalene muscles approximately 1 cm apart. The (C) &#34;active&#34; electrode (E1) and (D) &#34;reference&#34; recording electrode (E2) are positioned over the midclavicular line inferior to the last rib border, following the angle between the xiphoid process and the last costosternal cartilage. Additionally, (E) a disposable disk electrode is situated on the tail as a ground to minimize artifacts. Created with BioRender.com.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66568/66568fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66568/66568fig02.jpg" /> 
Figure 2: Compound muscle action potential. An illustration of a representative CMAP response recorded from the left hemidiaphragm muscle. CMAP peak to-peak amplitude is determined from negative peak voltage to positive peak voltage. Abbreviation: CMAP = Compound muscle action potential. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66568/66568fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66568/66568fig03.jpg" /> 
Figure 3: Two representative CMAP and MUNE recordings from a control rat, and an adult rat seven days after CTB-SAP intrapleural injection. (A) Left phrenic compound muscle action potential (CMAP) in a control adult rat (12 weeks of age) with peak-to-peak amplitude of 14.5 mV. Screen sensitivity = 1 mV/division and screen duration 1 ms/division. (B) Ten subsequent incremental responses with a total amplitude of 1.940 mV are divided by 10 to determine the average SMUP size (0.194 mV). Screen sensitivity = 0.2 mV/division and sweep speed of 1 ms/division. Calculated MUNE = 74 (MUNE=CMAP/average SMUP (14.5 mV/ 0.194 mV)) (C) Phrenic CMAP seven days following CTB-SAP injection showing almost no change in CMAP peak-to-peak amplitude 14.25 mV. Screen sensitivity = 1 mV/division and sweep speed of 1 ms/division. (D) Ten subsequent incremental responses with a total peak-to-peak amplitude of 2.360 mV divided by 10 to obtain an average SMUP size of 0.236 mV. Screen sensitivity = 0.2 mV/division and a sweep speed of 1 ms/division. Thus, the calculated MUNE was 60. Abbreviations: CMAP = compound muscle action potential; MUNE = motor unit number estimation; CTB-SAP = cholera toxin B fragment conjugated to saporin; SMUP = single motor unit potential. *Illustrates the artifact caused by electrical stimulation; it is important to note that due to the contrast between the amplitude of the electrical stimulus and CMAP, as well as incremental responses, we graphed only a portion of the stimulus. Blind-colored friendly colors have been utilized to enhance the visualization of the incremental responses for improved accessibility. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66568/66568fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66568/66568fig04.jpg" /> 
Figure 4: Electrophysiological assessment of CMAP, SMUP, and MUNE following CTB-SAP. No baseline differences between groups were found (seven 11-week-old male rats in both CTB-SAP and control groups). (A) CMAP showed no significant change. (B)&#160;The average SMUP of CTB-SAP rats showed a significant change over time (p &#60; 0.05) and with CTB-SAP (p &#60; 0.01), with no significant interaction, indicating a 50-60% increase in SMUP amplitude. (C) Seven days after CTB-SAP injection, the mean MUNE significantly decreased by 27.07%, from 102.58 &#177; 20.54 to 72.43 &#177; 15.59 estimated functional motor units (Mean &#177; SD). This reduction persisted, with an additional 11.75% decline at 28 days (MUNE: 60.64 &#177; 11.33, p &#60; 0.01). All measurements at baseline, 7, 14, and 28 days after injection were obtained from 14 rats, with n = 7 rats injected with conjugated CTB-SAP and n = 7 rats receiving unconjugated CTB and SAP as the control group. Six rats were used on day 21. CMAP, SMUP, and MUNE data are presented as mean, and comparisons were performed using two-way ANOVA. Asterisks indicate differences at various time points: *p &#60; 0.05, and **p &#60; 0.01. Abbreviations: CMAP = compound muscle action potential; MUNE = motor unit number estimation; CTB-SAP = cholera toxin B fragment conjugated to saporin; SMUP = single motor unit potential. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66568/66568fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Read this Scientific Article Protocol about Assessing Rat Diaphragm Motor Unit Connectivity Outcome Measures as Quantitative Biomarkers of Phrenic Motor Neuron Degeneration and Compensation at JoVE.co

Author Spotlight: Studying Neuromuscular Responses and Motor Neuron Plasticity in Neurodegenerative Diseases

In this study, we present an in vivo method for estimating motor unit number and size to quantify rat diaphragm motor unit connectivity. A step-by-step approach to these techniques is described.

Assessing Rat Diaphragm Motor Unit Connectivity Outcome Measures as Quantitative Biomarkers of Phrenic Motor Neuron Degeneration and Compensation

Loss of ventilatory muscle function is a consequence of motor neuron injury and neurodegeneration (e.g., cervical spinal cord injury and amyotrophic ...

assessing-rat-diaphragm-motor-unit-connectivity-outcome-measures-as

Nanyang Technological University

Research

JoVE Journal

Neuroscience

301 Views.  University of Missouri. We zijn geïnteresseerd in het begrijpen van normale en onaangepaste reacties van het neuromusculaire systeem in de context van gezondheid, veroudering en ziekte. Concreet willen we onderzoeken hoe het respiratoire motorische systeem reageert op verlies van motorneuronen door middel van compensatiemechanismen zoals collateral sprouting. Momenteel zijn verschillende arbeidstechnieken, zoals retrograde tracers, adenovirussen en immunohistochemie, gebruikt om he...

Video: Auteur in de schijnwerpers: onderzoek naar neuromusculaire reacties en plasticiteit van motorneuronen bij neurodegeneratieve ziekten

Paired Patch Clamp Recordings from Motor-neuron and Target Skeletal Muscle in Zebrafish

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle. This is a direct consequence of the accessibility to both cell types and ability to visually distinguish the single segmental CaP motor-neuron on the basis of morphology and location. This video demonstrates the microscopic methods used to identify a CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type. Identification of the CaP motor-neuron type is confirmed by either dye filling or by the biophysical features such as action potential waveform and cell input resistance. Motor-neuron recordings routinely last for one hour permitting long-term recordings from multiple different target muscle cells. Control over the motor-neuron firing pattern enables measurements of the frequency-dependence of synaptic transmission at the neuromuscular junction. Owing to a large quantal size and the low noise provided by whole cell voltage clamp, all of the unitary events can be resolved in muscle. This feature permits study of basic synaptic properties such as release properties, vesicle recycling, as well as synaptic depression and facilitation. The advantages offered by this in vivo preparation eclipse previous neuromuscular model systems studied wherein the motor-neurons are usually stimulated by extracellular electrodes and the muscles are too large for whole cell patch clamp. The zebrafish preparation is amenable to combining electrophysiological analysis with a wide range of approaches including transgenic lines, morpholino knockdown, pharmacological intervention and in vivo imaging. These approaches, coupled with the growing number of neuromuscular disease models provided by mutant lines of zebrafish, open the door for new understanding of human neuromuscular disorders.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target skeletal muscle. This video demonstrates the microscopic methods used to identify a segmental CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle.  ...

A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

The Neuromuscular Junction: Measuring Synapse Size, Fragmentation and Changes in Synaptic Protein Density Using Confocal Fluorescence Microscopy

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. Structural changes at the NMJ can result in neurotransmission failure, resulting in weakness, atrophy and even death of the muscle fiber. Many studies have investigated how genetic modifications or disease can alter the structure of the mouse NMJ. Unfortunately, it can be difficult to directly compare findings from these studies because they often employed different parameters and analytical methods. Three protocols are described here. The first uses maximum intensity projection confocal images to measure the area of acetylcholine receptor (AChR)-rich postsynaptic membrane domains at the endplate and the area of synaptic vesicle staining in the overlying presynaptic nerve terminal. The second protocol compares the relative intensities of immunostaining for synaptic proteins in the postsynaptic membrane. The third protocol uses Fluorescence Resonance Energy Transfer (FRET) to detect changes in the packing of postsynaptic AChRs at the endplate. The protocols have been developed and refined over a series of studies. Factors that influence the quality and consistency of results are discussed and normative data are provided for NMJs in healthy young adult mice.

The neuromuscular junction (NMJ) is altered in a variety of conditions that can sometimes culminate in synaptic failure. This report describes fluorescence microscope-based methods to quantify such structural changes.

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. ...

Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and morphological levels. Compound muscle action potential (CMAP) recording following phrenic nerve stimulation can be used to quantitatively assess functional diaphragm innervation by PhMNs of the cervical spinal cord in vivo in anesthetized rats and mice. Because CMAPs represent simultaneous recording of all myofibers of the whole hemi-diaphragm, it is useful to also examine the phenotypes of individual motor axons and myofibers at the diaphragm NMJ in order to track disease- and therapy-relevant morphological changes such as partial and complete denervation, regenerative sprouting and reinnervation. This can be accomplished via whole-mount immunohistochemistry (IHC) of the diaphragm, followed by detailed morphological assessment of individual NMJs throughout the muscle. Combining CMAPs and NMJ analysis provides a powerful approach for quantitatively studying diaphragmatic innervation in rodent models of CNS and PNS disease.

Compound muscle action potential recording quantitatively assesses functional diaphragm innervation by phrenic motor neurons. Whole-mount diaphragm immunohistochemistry assesses morphological innervation at individual neuromuscular junctions. The goal of this protocol is to demonstrate how these two powerful methodologies can be used in various rodent models of spinal cord disease.

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and ...

Measuring Neuromuscular Junction Functionality

Neuromuscular junction (NMJ) functionality plays a pivotal role when studying diseases in which the communication between motor neuron and muscle is impaired, such as aging and amyotrophic lateral sclerosis (ALS). Here we describe an experimental protocol that can be used to measure NMJ functionality by combining two types of electrical stimulation: direct muscle membrane stimulation and the stimulation through the nerve. The comparison of the muscle response to these two different stimulations can help to define, at the functional level, potential alterations in the NMJ that lead to functional decline in muscle.
Ex vivo preparations are suited to well-controlled studies. Here we describe an intensive protocol to measure several parameters of muscle and NMJ functionality for the soleus-sciatic nerve preparation and for the diaphragm-phrenic nerve preparation. The protocol lasts approximately 60 min and is conducted uninterruptedly by means of a custom-made software that measures the twitch kinetics properties, the force-frequency relationship for both muscle and nerve stimulations, and two parameters specific to NMJ functionality, i.e. neurotransmission failure and intratetanic fatigue. This methodology was used to detect damages in soleus and diaphragm muscle-nerve preparations by using SOD1G93A transgenic mouse, an experimental model of ALS that ubiquitously overexpresses the mutant antioxidant enzyme superoxide dismutase 1 (SOD1).

A functional assessment of the neuromuscular junction (NMJ) can provide essential information on the communication between muscle and nerve. Here we describe a protocol to comprehensively evaluate both the NMJ and muscle functionality using two different muscle-nerve preparations, i.e. soleus-sciatic and diaphragm-phrenic.

Neuromuscular junction (NMJ) functionality plays a pivotal role when studying diseases in which the communication between motor neuron and muscle is ...

A Simple Neuronal Mechanical Injury Methodology to Study Drosophila Motor Neuron Degeneration

The degeneration of neurons occurs during normal development and in response to injury, stress, and disease. The cellular hallmarks of neuronal degeneration are remarkably similar in humans and invertebrates as are the molecular mechanisms that drive these processes. The fruit fly, Drosophila melanogaster, provides a powerful yet simple genetic model organism to study the cellular complexities of neurodegenerative diseases. In fact, approximately 70% of disease-associated human genes have a Drosophila homolog and a plethora of tools and assays have been described using flies to study human neurodegenerative diseases. More specifically the neuromuscular junction (NMJ) in Drosophila has proven to be an effective system to study neuromuscular diseases because of the ability to analyze the structural connections between the neuron and the muscle. Here, we report on an in vivo motor neuron injury assay in Drosophila, which reproducibly induces neurodegeneration at the NMJ by 24 h. Using this methodology, we have described a temporal sequence of cellular events resulting in motor neuron degeneration. The injury method has diverse applications and has also been utilized to identify specific genes required for neurodegeneration and to dissect transcriptional responses to neuronal injury.

A Simple Neuronal Mechanical Injury Methodology to Study Drosophila Motor Neuron Degeneration

Here we describe a simple and widely accessible method to injure segmental nerves in Drosophila larvae to visualize and quantify neurodegeneration of motor neurons at the neuromuscular junction (NMJ) of third instar larvae.

The degeneration of neurons occurs during normal development and in response to injury, stress, and disease. The cellular hallmarks of neuronal ...

Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated measurements over weeks or months of accessory respiratory muscle activity while simultaneously measuring ventilation in a non-anesthetized, freely behaving mouse. The technique includes the surgical implantation of a radio transmitter and the insertion of electrode leads into the scalene and trapezius muscles to measure the electromyogram activity of these inspiratory muscles. Ventilation is measured by whole-body plethysmography, and animal movement is assessed by video and is synchronized with electromyogram activity. Measurements of muscle activity and ventilation in a mouse model of amyotrophic lateral sclerosis are presented to show how this tool can be used to investigate how respiratory muscle activity changes over time and to assess the impact of muscle activity on ventilation. The described methods can easily be adapted to measure the activity of other muscles or to assess accessory respiratory muscle activity in additional mouse models of disease or injury.

This paper introduces a method for repeated measurements of ventilation and respiratory muscle activity in a freely behaving amyotrophic lateral sclerosis (ALS) mouse model throughout disease progression with whole-body plethysmography and electromyography via an implanted telemetry device.

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated ...

Medicine

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...

Standardized Evaluation of DIAm EMG Post-C2SH in Rats to Minimize Variability

Assessing Functional Recovery of Eupneic Diaphragm Activity Following Unilateral Cervical Spinal Cord Hemisection in Rats

Following cSCI, activation of the DIAm can be impacted depending on the extent of the injury. The present manuscript describes a unilateral C2 hemisection (C2SH) model of cSCI that disrupts eupneic ipsilateral diaphragm (iDIAm) electromyographic (EMG) activity during breathing in rats. To evaluate recovery of DIAm motor control, the extent of deficit due to C2SH must first be clearly established. By verifying a complete initial loss of iDIAm EMG during breathing, subsequent recovery can be classified as either absent or present, and the extent of recovery can be estimated using the EMG amplitude. Additionally, by measuring the continued absence of iDIAm EMG activity during breathing after the acute spinal shock period following C2SH, the success of the initial C2SH may be validated. Measuring contralateral diaphragm (cDIAm) EMG activity can provide information about the compensatory effects of C2SH, which also reflects neuroplasticity. Moreover, DIAm EMG recordings from awake animals can provide vital physiological information about the motor control of the DIAm after C2SH. This article describes a method for a rigorous, reproducible, and reliable C2SH model of cSCI in rats, which is an excellent platform for studying respiratory neuroplasticity, compensatory cDIAm activity, and therapeutic strategies and pharmaceuticals.

Author Spotlight: Neuromotor Control and Recovery of Diaphragm Function Following Cervical Spinal Hemisection in Rats

Respiratory complications are the leading cause of death in individuals with cervical spinal cord injury (cSCI). Animal models of cSCI are essential for mechanistic evaluations and pre-clinical studies. Here, we introduce a reproducible method to assess functional recovery of diaphragm muscle (DIAm) activity following unilateral C2 spinal hemisection (C2SH) in rats.

Following cSCI, activation of the DIAm can be impacted depending on the extent of the injury. The present manuscript describes a unilateral C2 hemisection ...

Measuring Compound Muscle Action Potentials by Stimulating the Phrenic Nerve in a Rat

Source: Martin, M. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/52605">Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons.</a> J. Vis. Exp. (2015)
This video describes recording compound muscle action potentials (CMAPs) in diaphragm muscle fibers in a rat through phrenic nerve stimulation. A high-intensity electric pulse stimulation of the phrenic nerve generates action potentials in the diaphragm muscle fibers, which are observed by obtaining the response curve.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Assessing Rat Diaphragm Motor Unit Connectivity Outcome Measures as Quantitative Biomarkers of Phrenic Motor Neuron Degeneration and Compensation

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