Name: functional and morphological assessment of diaphragm innervation by phrenic motor neurons
Uploaded: 2015-05-25T14:00:00.000+00:00
Duration: 9 min 43 s
Description: Watch this Scientific Journal Video about Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons at JoVE.com

This work was supported by the NINDS (grant #1R01NS079702 to A.C.L.) and the SURP Program at Thomas Jefferson University (M.M.).

Experimental procedures were approved by the Thomas Jefferson University institutional animal care and use committee and conducted in compliance with the European Communities Council Directive (2010/63/EU, 86/609/EEC and 87-848/EEC), the NIH Guide for the care and use of laboratory animals, and the Society for Neuroscience&#8217;s Policies on the Use of Animals in Neuroscience Research.
1. Compound Muscle Action Potentials (CMAPs)
<ol>
	<li>Preparing the animal:
	<ol>
		<li>Anesthetize the rat using an inhalant (isoflurane) at 1-2% delivered to effect or by injection (ketamine, xylazine, acepromazine). Dosage of injectable anesthesia is as follows: 95.0 mg/kg of ketamine, 10.0 mg/kg of xylazine, 0.075 mg/kg of acepromazine. 
		NOTE: We successfully use both inhalant and injectable anesthetic approaches when conducting this CMAP recording protocol, with no preference.</li>
		<li>The animal must be maintained at the appropriate anesthetic depth before the surgery is started, through the conclusion of surgery, and until post-operative buprenorphine has taken effect. Confirm proper anesthetization by brief toe pinch to determine that a withdrawal response is not elicited. Additionally, test the corneal reflex with a cotton swab. Throughout the surgical procedure, determine that heart rate and respiratory rate have not increased, which suggests that anesthetic depth has become too light.</li>
		<li>Apply a vet ointment on the animal&#8217;s eyes to prevent dryness while under anesthesia.</li>
		<li>The surgeon should wash her/his hands with a disinfectant (chlorhexidine scrub) before beginning surgery. The surgeon should wear sterile gloves, facemask, and gown or clean lab coat. All instruments should be washed and autoclaved before surgery. Add a sterilization indicator strip inside the instrument kit before autoclaving at the level of the instruments to verify that sterilization is complete. The surgeon should verify that the entire line on the strip is black before beginning surgery. Tape the outside of the box with indicating autoclave tape as well. If surgery is to be performed on several animals on the same day, clean the instruments between surgeries and sterilize in a glass bead sterilizer. If necessary, sterilize both ends of the instrument in the glass bead sterilizer by placing the handle of the instrument in the sterilizer for the appropriate time, removing the instrument with a sterile hemostat, cooling it in a sterile field and then placing the functional end of the instrument into the glass bead sterilizer for the appropriate period of time.</li>
		<li>Once animal is anesthetized, place on surgical board with dorsal surface down and abdomen facing up so that tape can be used to secure animal&#8217;s forelimbs to surgical board.</li>
		<li>Shave ventral surface caudally beginning at the base of the skull, and ensure that the area around abdomen on either side of the midline is shaved depending on which hemi-diaphragm is being recorded. Apply povidone-iodine to the shaved surface of the skin.</li>
	</ol>
	</li>
	<li>Preparation for CMAP recordings:
	<ol>
		<li>Following animal preparation, place ground electrode subcutaneously into tail (Figure 1).</li>
		<li>Place the reference electrode subcutaneously into the contralateral lower abdominal region.</li>
		<li>Place conductive gel onto self-adhesive surface strip or disk recording electrode (dimensions of surface strip electrode: 0.5 x 3 cm), and then place surface electrode transversely along the unilateral costal margin.</li>
		<li>Place stimulating electrodes transcutaneously 0.5 cm apart lateral to trachea and superior to the clavicle. Insert the electrodes approximately 1.0 cm deep through the skin. Secure the electrodes by hand so that their location does not move with subsequent stimulations. 
		NOTE: Angle stimulating electrodes 90&#176; relative to the skin at the site of insertion.</li>
	</ol>
	</li>
	<li>Compound Muscle Action Potential (CMAP) recordings:
	<ol>
		<li>Obtain electrophysiological recordings using a stimulator/amplifier followed by computer assisted data analysis.</li>
		<li>The stimulus parameters of single pull stimulation should be 0.5 msec, 1.0 Hz supramaximal pulses. The amplitude of the stimulus should be between 6.0 and 8.0 V (Figure 2A). 
		NOTE: Within an experiment, use a stimulus intensity with the same amplitude across animals. The objective is to maximize the response amplitude by using the minimal amplitude of stimulation. It is important to note that the initial response that occurs at the time of stimulation is a stimulation artifact. In addition, care should be taken to obtain a CMAP response that is preceded by a stable/flat baseline after the stimulation artifact, which is then followed by the rapid CMAP response. In order to obtain such a response, it may be necessary to reposition the stimulation and/or recording electrodes.</li>
		<li>Wait 30 sec between stimulations, and repeat 10 stimulations in a row to obtain the average response (Figure 2B-C). Measure the amplitude from baseline to peak (Figure 2A). Conduct phrenic nerve stimulation during the phase of the animal&#8217;s respiratory cycle when the phrenic nerve/diaphragm is not activated. The procedure can be repeated on the same animal for the contralateral hemi-diaphragm.</li>
		<li>After the final CMAP recording session, perfuse the animal if NMJ staining will be conducted. Conduct CMAP recordings repeatedly on the same animal approximately once every week (or at interval of choice).</li>
		<li>Following survival CMAP recordings, allow animal to recover on a circulating warm water heating pad, and continuously monitor during recovery until awake and sternally recumbent. Do not return an animal that has undergone surgery to the company of other animals until fully recovered.</li>
		<li>Once sternally recumbent, monitor the animal every 12 hr for the first 48 hr (once daily thereafter). If the animal appears to be dehydrated (flaccid skin, listlessness), administer fluids (lactated ringers solution; subcutaneously; 1-2 ml per injection, with the sites rotated; 2 times per day) until post-surgical weight and fluid loss stabilizes. Provide animals with softened food in small dishes during the acute post-operative period if necessary to encourage eating if they cannot reach the wire bar lid feeder.</li>
		<li>Administer buprenorphine at a dose of 0.05 mg/kg before the end of surgery and then at 12 hr intervals for the first 24 hr (and dependent upon signs of pain/stress thereafter) via subcutaneous injection (site rotated). If signs of pain or distress are noticed thereafter, treat animals with buprenorphine. Signs indicative of pain/distress include lack of activity, vocalizations, lack of eating or drinking (dehydration), excessive weight loss, guarding, excessive gnawing or scratching at site of incision. Additional criteria used in determining pain and distress include unresponsiveness to extraneous stimuli, transient or unprovoked vocalization, labored or abnormal breathing, persistent reddish-brown naso-occular discharge, marked piloerection. Also monitor evidence of poor hygiene that would suggest ill animals. If animal is still observed to be in distress after 72 hr of treatment (according to the pain/stress criteria outline above, and after receiving the analgesic protocol described above) euthanize the animal.</li>
	</ol>
	</li>
</ol>
2. Diaphragm Neuromuscular Junction (NMJ) Analysis
<ol>
	<li>Animal dissection:
	<ol>
		<li>Administer the animal an overdose of injectable anesthesia (3 times dose used above: ketamine at 285.0 mg/kg, xylazine at 30.0 mg/kg, acepromazine at 0.225 mg/kg; administered intraperitoneally). Following overdose, all animals should have a second method of euthanasia (thoracotomy) to ensure death. Death should be confirmed by observing cardiac and respiratory arrest and by noting the animal&#39;s fixed and dilated pupils.</li>
		<li>Conduct a laparotomy with scalpel or dissection scissors extending the excision from the zyphoid process caudally along the midline. Carefully separate skin and connective tissue from underlying muscle using scalpel and forceps.</li>
		<li>While pulling up on the zyphoid process, use dissection scissors to remove whole diaphragm, ensuring that the diaphragm remains attached to surrounding bone. This is important as it will prevent damage to the diaphragm muscle, will allow for successful dissection of the entire diaphragm and will aid in cleaning muscle for subsequent staining. 
		NOTE: At this point, animal can be perfused if necessary.</li>
	</ol>
	</li>
	<li>Cleaning and staining of the diaphragm:
	<ol>
		<li>Pin down the whole dissected diaphragm, superior surface facing up, to silicone rubber in a glass 100 mm Petri dish in 4% paraformaldehyde (made in 1% phosphate buffered saline - PBS) for 20 min using a stereomicroscope. Stretch the diaphragm to make it taut, and using insect pins, pin down the surrounding tissue to avoid pinning the diaphragm muscle itself (Figure 3).</li>
		<li>Carefully clean off connective tissue from only the superior surface of muscle with #5 forceps and small scissors. 
		NOTE: Carry out all subsequent steps of this staining protocol slowly rocking at room temperature covered from light unless otherwise specified. Ensure that the diaphragm muscle is always completely submerged in liquid to avoid drying out.</li>
		<li>Prior to staining, prepare all necessary reagents: 1x PBS, 2% bovine serum albumin (BSA)/PBS, 0.1 M glycine made in 2% BSA/PBS, 4% paraformaldehyde (PFA) in 1x PBS, and 0.2% TBP (0.2% Triton X100 in 2% BSA/PBS). Be sure to pre-cool methanol at -20 &#176;C.</li>
		<li>Wash 3x for 10 min in 1x PBS.</li>
		<li>Incubate in 0.1 M glycine (made in 2% BSA/PBS) for 30 min.</li>
		<li>Incubate in RBTX (rhodamine-conjugated &#945;-bungarotoxin) solution for 15 min. Solution for RBTX is as follows: Rhodamine-conjugated alpha bungarotoxin 1:400 dilution in 1x PBS. 
		NOTE: In this protocol, Rhodamine is used, but any fluorophore of choice can be used for staining.</li>
		<li>Wash 3x for 10 min in 1x PBS.</li>
		<li>In -20 &#176;C freezer, incubate with MeOH for 5 min. 
		NOTE: Do not add MeOH before placing muscle in the freezer, and immediately remove solution following the incubation period.</li>
		<li>Wash 3x for 10 min in 1x PBS.</li>
		<li>Block in 0.2% TBP for 1 hr (0.2% Triton made in 2% BSA/PBS).</li>
		<li>Incubate with primary antibody solution (in 0.2% TBP) at 4 &#176;C overnight while rocking. Dilutions of antibodies as follows: SV2 (1:10) and SMI-312 (1:1,000).</li>
		<li>Wash 3x for 10 min with 0.2% TBP.</li>
		<li>Incubate with secondary antibody using 1:100 dilution of FGM1 (FITC-conjugated goat anti-mouse IgG1; made in 0.2% TBP) while rocking for 1 hr. 
		NOTE: Protect from light for the rest of the protocol for staining.</li>
		<li>Wash 3x for 10 min with 1x PBS.</li>
		<li>Re-clean surrounding connective tissue on the superior (see above) surface of muscle prior to mounting and coverslipping. 
		NOTE: It is recommended to clean and then mount sample immediately after staining. However, if necessary, sample may be covered in 1x PBS, wrapped in parafilm to prevent evaporation and stored at 4 &#176;C for up to 1 week, with changing of PBS daily.</li>
		<li>Upon mounting on glass slide, coverslip with non-hardset Vectashield. Seal edges of coverslip with clear nail polish to prevent drying out. Store slides lying flat in cardboard slide folders in -20 &#176;C for long term.</li>
	</ol>
	</li>
	<li>Analysis and Confocal Imaging:
	<ol>
		<li>Quantitatively analyze the morphology of the stained and mounted hemi-diaphragm under 20X and 40X magnification using an upright epifluorescence microscope. 
		NOTE: Because the diaphragm is stained and analyzed in a whole-mount fashion, the muscle is quite thick. Consequently, most of the fluorescence staining is completely out of focus when a static image is taken using non-confocal fluorescence microscopy. For this reason, it is best to conduct the NMJ analysis in real-time on the microscope as this allows you to constantly focus &#8220;up&#8221; and &#8220;down&#8221; through the muscle to properly identify the morphology of each of the individual NMJs. Using this approach, one can easily follow the trajectory of individual motor axons as they move through the tissue in spatial relationship to the post-synaptic receptor clusters.</li>
		<li>By measuring the ventral-to-dorsal length of the muscle, divide the muscle into 3 separate sections for analysis based on topographic innervation patterns from specific spinal cord levels (Figure 4H).</li>
		<li>Beginning at the ventral region of the muscle and moving dorsally, analyze all en face NMJs located on surface muscle fibers. Do not analyze muscles fibers deeper than the first 3-4 fibers from the surface as interpretation may be difficult due to poorer antibody penetration (RBTX penetrates much deeper into the muscle due to the small size of the toxin compared to the larger antibodies used to label neurons).</li>
		<li>Identify each NMJ as intact if the pre-terminal axon of the neuron is thick in diameter and all regions of the nerve terminal completely overlap with the underlying muscle acetylcholine receptors (AChRs).</li>
		<li>Identify each NMJ as partially denervated if any region of the underlying muscle AChRs is unoccupied with corresponding nerve terminal.</li>
		<li>Identify each NMJ as completely denervated if the muscle AChRs have no corresponding nerve terminal (it is also important to have other NMJs with labeled neurons in the same field of focus, so that one can be sure the lack of nerve terminal is not simply due to poor antibody penetration in that region of muscle).</li>
		<li>Identify each NMJ as multiply innervated if there is more than one pre-terminal axon innervating an individual NMJ (this indicates that there has been some level of denervation and likely attempts of reinnervation at this junction).</li>
		<li>Identify each NMJ as thinly innervated if the NMJ has complete overlap between nerve terminal and underlying AChRs but has a thin pre-terminal axon (this also is indicative of some level of denervation and attempts at reinnervation).</li>
		<li>Conduct analysis for each of the 3 identified regions for all of the above NMJ categories for each animal. Average the data from each animal with results from other animals from the same experimental group. Approximately 200-300 junctions in rat hemi-diaphragm muscle and 150-200 in mouse hemi-diaphragm should be quantified in at least 3 (preferably 5-6 or more) animals per experimental group.</li>
		<li>Obtain high-resolution confocal images, used for publication, with a confocal microscope.</li>
		<li>Obtain Z-stacks at 0.3 &#956;m step size for 20-40 &#956;m depths, and collect additional optical sections above and below each junction to ensure that the entire synaptic profile is included.</li>
		<li>Use acquisition software to reconstruct z-series images into maximum intensity projections.</li>
	</ol>
	</li>
</ol>

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We have nothing to disclose.

As respiratory function is compromised in both traumatic SCI and ALS, developing therapies that target breathing and specifically diaphragm innervation are clinically relevant5,6. In order to comprehensively study respiratory function, a combined approach method should be used. CMAPs measure the degree of functional innervation of the diaphragm by way of external phrenic nerve stimulation, but not endogenous bulbospinal respiratory drive8. In addition, these recordings do not allow for examination of morphological changes at the NMJ, particularly at the level of individual motor axons and post-synaptic acetylcholine receptors. By using the described staining protocol, one can quantitatively assess relevant morphological phenotypes such as complete denervation, partial denervation and even regenerative sprouting and reinnervation at individual NMJs of the diaphragm muscle9.
Nevertheless, these two protocols still do not fully capture all aspects of respiratory pathology. For example, if injury occurs to the descending bulbospinal axons without any damage to the PhMN pool or phrenic nerve, CMAP amplitudes may still appear normal even though there is significant impairment in this circuitry because the phrenic nerve is externally stimulated14. Importantly, CMAPs do not allow for measurement of endogenous supraspinal respiratory drive. Furthermore, CMAPs do not provide a measure of overall ventilation. It may be beneficial to conduct, for example, whole body plethysmography to examine overall breathing behavior and spontaneous intra-muscular EMG recordings and phrenic nerve recordings to test endogenous PhMN activation. In addition, other approaches such as measuring blood gas levels can provide important information about respiratory function15. It may also be possible to employ the described diaphragm CMAP recording technique for motor unit number estimation16, though we have never attempted this. It is also important to note that these two protocols were described for only one respiratory muscle, the diaphragm. However, there are other inspiratory and expiratory respiratory muscles such as the intercostals that can be studied and that are affected by diseases such as SCI and ALS.
In this protocol, we describe diaphragm CMAP and NMJ analysis specifically in the rat model. Similarities between rats and mice make transfer of both these techniques to mice straightforward. We previously showed that partial loss of PhMNs following cervical contusion SCI does result in quantifiable decreases in diaphragm CMAP amplitude that correlate with morphological NMJ denervation. Furthermore, we have demonstrated these deficits in both mouse17 and rat10-12 SCI models. In addition, we have shown quantifiable diaphragm CMAP reduction in both mouse18 and rat13,19 models of ALS (i.e., the SOD1G93A model) that are associated with more subtle diaphragm denervation by PhMNs compared to cervical SCI, and we have demonstrated that we can detect relatively smaller therapeutic effects on CMAP amplitude of interventions such as stem cell transplantation in these ALS models19. Collectively, these findings demonstrate the utility of using these analyses to quantitatively assess both functional and morphological assessment of diaphragm innervation by PhMNs in both rat and mouse models of SCI and ALS.
By combining the histological and functional techniques detailed in this protocol to examine breathing and innervation at the NMJ, better insight can be gained into nervous system diseases associated with diaphragm dysfunction, thus providing greater awareness of how to target therapies in animal models and ultimately in patients.

Amyotrophic Lateral Sclerosis (ALS) is a debilitating motor neuron disease associated with the loss of both upper and lower motor neurons and consequent muscle paralysis. Upon diagnosis, patient survival is on average only 2-5 years1. Phrenic motor neuron (PhMN) loss is a critical component of the pathogenesis of ALS. Patients ultimately die due to loss of PhMN innervation of the diaphragm, the primary muscle of inspiration2,3. Traumatic spinal cord injury (SCI) is also a serious problem with associated breathing difficulties. Approximately 12,000 new cases of SCI occur each year4 due to traumatic damage to the spinal cord. Despite disease heterogeneity with respect to location, type and severity, the majority of SCI cases involve trauma to the cervical spinal cord, which often results in debilitating and persistent respiratory compromise. In addition to ALS and SCI, other central nervous system (CNS) diseases can be associated with diaphragmatic respiratory dysfunction5,6.
The phrenic nerve is an efferent motor nerve that innervates the ipsilateral hemi-diaphragm and that originates from PhMN cell bodies located in the C3-C5 levels of the ipsilateral cervical spinal cord. PhMN output is controlled by descending bulbospinal input from the brainstem in an area known as the rostral ventral respiratory group (rVRG)7. The rVRG-PhMN-diaphragm circuit is central to the control of inspiratory breathing, as well as other non-ventilatory diaphragm behaviors. Various traumatic injuries and neurodegenerative disorders that affect this circuitry can lead to a profound decline in respiratory function and patient quality of life. Descending input to PhMNs from the rVRG, PhMN survival, phrenic nerve integrity and proper innervation at the diaphragm neuromuscular junction (NMJ) are all necessary for normal diaphragm function. It is therefore important to employ techniques that can quantitatively evaluate this circuit in vivo in rodent models of ALS, SCI and other CNS diseases.
With this protocol, the goal is to describe experimental tools for assessing PhMN innervation of the diaphragm at both the electrophysiological and morphological levels. Compound muscle action potentials (CMAPs) are recorded by stimulating all efferent motor neuron axons of a given motor nerve and then analyzing the elicited depolarization response of the target myofibers. This technique can be used in vivo in anesthetized rats and mice to quantify functional innervation of the hemi-diaphragm by PhMNs8. Due to the fact that CMAPs represent simultaneous recording of all (or at least many/most) 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 muscle9. Combining CMAPs and NMJ analysis provides a powerful approach for quantitatively studying diaphragmatic innervation in rodent models of CNS and PNS disease.

<table><tbody><tr><td>Paraformaldehyde</td><td>Fisher</td><td>T353-500</td><td>Make 10% solution first in de-ionized distilled water; make 4% with 1x PBS, adjust pH to 7.4</td></tr><tr><td>1x Phosphate buffered saline, pH 7.4</td><td>Invitrogen</td><td>10010049</td><td></td></tr><tr><td>2% Bovine serum albumin (2% BSA)</td><td>Sigma-Aldrich</td><td>A3059-100g</td><td>Dissolve 2 g BSA into 100 ml of 1x PBS</td></tr><tr><td>0.2% Triton X100 in 2% BSA/PBS (blocking buffer)</td><td>Sigma-Aldrich</td><td>T9284-100mL</td><td>Dissolve 0.2 ml/100 ml 2% BSA/PBS</td></tr><tr><td>0.1 M Glycine</td><td>Sigma-Aldrich</td><td>G-7126</td><td>Add 0.185 g to 25 ml of 2% BSA/PBS</td></tr><tr><td>&amp;alpha;-bungarotoxin</td><td>Invitrogen</td><td>T1175</td><td>Concentration 1:400</td></tr><tr><td>SMI-312</td><td>Sternberger Monoclonals</td><td>SMI312</td><td>Concentration 1:1,000</td></tr><tr><td>SV2</td><td>Developmental Studies Hybridoma Bank</td><td>SV2-Supernatant</td><td>Concentration 1:10</td></tr><tr><td>FITC goat anti-mouse IgG1</td><td>Roche</td><td>3117731001</td><td>Concentration 1:100</td></tr><tr><td>Silicone rubber</td><td>Sylgard, Dow Corning</td><td>Part # 184</td><td>Follow instructions that come with kit: can use multiple sized culture dish (30 mm, 60 mm, 100 mm) depending on needs</td></tr><tr><td>Vectashield fluorescent mounting medium</td><td>Vector laboratories</td><td>H-1000</td><td>This is not a hard-set medium. You will need to secure the cover slip with clear nail polish.</td></tr><tr><td>Small spring scissors</td><td>Fine Science Tools</td><td>15002-08</td><td></td></tr><tr><td>Dissection forceps</td><td>Fine Science Tools</td><td>11295-51</td><td></td></tr><tr><td>Software for CMAP recordings</td><td>Scope 3.5.6; ADI</td><td></td><td></td></tr><tr><td>Disk surface electrodes</td><td>Natus neurology</td><td>019-409000</td><td></td></tr><tr><td>Subdermal needle electrodes</td><td>Natus neurology</td><td>019-453100</td><td></td></tr><tr><td>Conductive gel</td><td>Aquasonic</td><td>122-73720</td><td></td></tr><tr><td>Stimulator/recording system for CMAP recordings</td><td>ADI Powerlab 8SP stimulator</td><td></td><td></td></tr><tr><td>Amplifier for CMAP recordings</td><td>BioAMP</td><td></td><td></td></tr></tbody></table>

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.

Adult Sprague-Dawley rats received either laminectomy only (uninjured control) or unilateral hemi-contusion SCI at the C4 spinal cord level10-12. At 5 weeks post-surgery, peak CMAP amplitude recorded from the hemi-diaphragm ipsilateral to the laminectomy/injury site was significantly reduced in SCI rats (Figure 2C) compared to laminectomy-only control (Figure 2B). All NMJs in the hemi-diaphragm were completely intact in control non-diseased wild-type rats (Figure 4A,C). On the contrary, SOD1G93A rats (a rodent model of ALS) showed significant pathology at hemi-diaphragm NMJs (Figure 4B), including complete denervation (Figure 4G, arrowhead), partial denervation (Figure 4E, arrowhead; Figure 4G, NMJ on right), multiple innervation (Figure 4D) and thin pre-terminal axons (Figure 4F, arrowhead).
<img alt="Electrophysiology setup with electrode placements on a rat for nerve stimulation and recording." src="/files/ftp_upload/52605/52605fig1highres.jpg" width="400" data-altupdatedat="1747736610000" data-alt="Figure 1"> 
Figure 1: CMAP electrode placement. The ground electrode is placed subcutaneously into the tail. The reference electrode is placed into the abdomen. The surface recording electrode is placed transversely along the unilateral costal margin. Stimulating electrodes are depicted as both red and black. <a href="https://www.jove.com/files/ftp_upload/52605/52605fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Electrophysiology graph with mV vs. time, showing stimulation, baseline, peak amplitude." src="/files/ftp_upload/52605/52605fig2highres.jpg" width="650" data-altupdatedat="1747736610000" data-alt="Figure 2"> 
Figure 2: CMAP recordings and analysis. (A) The amplitude of CMAP response can be measured from baseline to peak. A typical CMAP response should show both the stimulus artifact and response curve. (B-C) Representative average CMAP responses. Stimulation should be carried out at least ten consecutive times to obtain a repeated average of the response. Shown here are a laminectomy-only uninjured rat (B) and a comparison of CMAP responses for an injured rat that received a unilateral C4 hemi-contusion (C). <a href="https://www.jove.com/files/ftp_upload/52605/52605fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Dissection setup: Frog dissection pinned specimen diagram in petri dish for anatomical study." src="/files/ftp_upload/52605/52605fig3highres.jpg" width="400" data-altupdatedat="1747736610000" data-alt="Figure 3"> 
Figure 3: Preparation of diaphragm for whole-mount staining. Insect pins should be placed into surrounding tissue to secure the hemi-diaphragm to Sylgard. Avoid placing pins into the muscle itself. In this image, both hemi-diaphragm muscles are shown and pinned down separately. <a href="https://www.jove.com/files/ftp_upload/52605/52605fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Fluorescence microscopy; SMI-32+SV2+alpha-bungarotoxin; neuromuscular junction analysis." src="/files/ftp_upload/52605/52605fig4highres.jpg" width="700" data-altupdatedat="1747736610000" data-alt="Figure 4"> 
Figure 4: NMJ phenotypes. Confocal imaging was conducted to visualize post-synaptic acetylcholine receptors (in red labeled with rhodamine-conjugated α bungarotoxin) and axonal fibers and pre-synaptic terminals (in green labeled with SMI-32 and SV2 antibodies, respectively). All NMJs in hemi-diaphragm were completely intact in control non-diseased wild-type rats (A, C). On the contrary, SOD1G93A rats (a rodent ALS model) showed significant pathology at hemi-diaphragm NMJs (B), including complete denervation (G, arrowhead), partial denervation (E, arrowhead; G, NMJ on right), multiple innervation (D) and thin pre-terminal axons (F, arrowhead). The diaphragm muscle is sub-divided into three separate sections for NMJ morphology analysis (H)13. Scale bar: 25 µm. <a href="https://www.jove.com/files/ftp_upload/52605/52605fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons at JoVE.com

Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

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 ...

functional-morphological-assessment-diaphragm-innervation-phrenic

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

17.5K Views.  Sidney Kimmel Medical College at Thomas Jefferson University. 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. 

Video: Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

Assessment of Motor Balance and Coordination in Mice using the Balance Beam

Brain injury, genetic manipulations, and pharmacological treatments can result in alterations of motor skills in mice. Fine motor coordination and balance can be assessed by the beam walking assay. The goal of this test is for the mouse to stay upright and walk across an elevated narrow beam to a safe platform. This test takes place over 3 consecutive days: 2 days of training and 1 day of testing. Performance on the beam is quantified by measuring the time it takes for the mouse to traverse the beam and the number of paw slips that occur in the process. Here we report the protocol used in our laboratory, and representative results from a cohort of C57BL/6 mice. This task is particularly useful for detecting subtle deficits in motor skills and balance that may not be detected by other motor tests, such as the Rotarod.

Deficits in fine motor coordination can be assessed with the balance beam test. Performance on the beam is quantified by the speed at which the beam is traversed and the number of times the mouse slips on the beam.

Brain injury, genetic manipulations, and pharmacological treatments can result in alterations of motor skills in mice.  Fine motor coordination and ...

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor functions. Most investigations of sensorimotor mechanisms rely on either examination of neurons while an animal is static1,2 or record extracellular neuronal activity during a movement.3,4 While these studies have provided the fundamental background for sensorimotor function, they either do not evaluate functional information which occurs during a movement or are limited in their ability to fully characterize the anatomy, physiology and neurochemical phenotype of the neuron. A technique is shown here which allows extensive characterization of individual neurons during an in vivo movement. This technique can be used not only to study primary afferent neurons but also to characterize motoneurons and sensorimotor interneurons. Initially the response of a single neuron is recorded using electrophysiological methods during various movements of the mandible followed by determination of the receptive field for the neuron. A neuronal tracer is then intracellularly injected into the neuron and the brain is processed so that the neuron can be visualized with light, electron or confocal microscopy (Fig. 1). The detailed morphology of the characterized neuron is then reconstructed so that neuronal morphology can be correlated with the physiological response of the neuron (Figs. 2,3). In this communication important key details and tips for successful implementation of this technique are provided. Valuable additional information can be determined for the neuron under study by combining this method with other techniques. Retrograde neuronal labeling can be used to determine neurons with which the labeled neuron synapses; thus allowing detailed determination of neuronal circuitry. Immunocytochemistry can be combined with this method to examine neurotransmitters within the labeled neuron and to determine the chemical phenotypes of neurons with which the labeled neuron synapses. The labeled neuron can also be processed for electron microscopy to determine the ultrastructural features and microcircuitry of the labeled neuron. Overall this technique is a powerful method to thoroughly characterize neurons during in vivo movement thus allowing substantial insight into the role of the neuron in sensorimotor function.

A technique is described to quantify the in vivo physiological response of mammalian neurons during movement and correlate the physiology of the neuron with neuronal morphology, neurochemical phenotype and synaptic microcircuitry.

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor ...

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.
This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath5, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.
To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation4,6,7. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.
We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g. in the suspended buccal mass preparation8 or in vivo9. This process can also be applied in other motor pools10,11,12 in Aplysia or in other animal systems2,3,13,14.

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

In animals with large  identified neurons (e.g. mollusks), analysis of motor pools is done using  intracellular techniques1,2,3,4. Recently,  we developed ...

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 ...

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 ...

Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea

Cochlear inner hair cells (IHCs) transmit acoustic signals to spiral ganglion neurons (SGNs) through ribbon synapses. Several experimental studies have indicated that hair cell synapses may be the initial targets in sensorineural hearing loss (SNHL). Such studies have proposed the concept of cochlear &#34;synaptopathy&#34;, which refers to alterations in ribbon synapse number, structure, or function that result in abnormal synaptic transmission between IHCs and SGNs. While cochlear synaptopathy is irreversible, it does not affect the hearing threshold. In noise-induced experimental models, restricted damage to IHC synapses in select frequency regions is employed to identify the environmental factors that specifically cause synaptopathy, as well as the physiological consequences of disturbing this inner ear circuit. Here, we present a protocol for analyzing cochlear synaptic morphology and function at a specific frequency region in adult mice. In this protocol, cochlear localization of specific frequency regions is performed using place-frequency maps in conjunction with cochleogram data, following which the morphological characteristics of ribbon synapses are evaluated via synaptic immunostaining. The functional status of ribbon synapses is then determined based on the amplitudes of auditory brainstem response (ABR) wave I. The present report demonstrates that this approach can be used to deepen our understanding of the pathogenesis and mechanisms of synaptic dysfunction in the cochlea, which may aid in the development of novel therapeutic interventions.

This manuscript describes an experimental protocol for evaluating the morphological characteristics and functional status of ribbon synapses in normal mice. The present model is also suitable for noise-induced and age-related cochlear synaptopathy-restricted models. The correlative results of previous mouse studies are also discussed.

Cochlear inner hair cells (IHCs) transmit acoustic signals to spiral ganglion neurons (SGNs) through ribbon synapses. Several experimental studies have ...

Quantifying Phrenic Motor Unit Function in Rodent Models of Neurodegeneration

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.

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.

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

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 ...

Diaphragmatic Ultrasound in Adults: Image Acquisition and Interpretation

Diaphragm dysfunction is a widely recognized concern across numerous medical specialties and clinical settings. Timely and accurate assessment of the diaphragm is vital not only in critically ill patients, where it has a role in weaning from mechanical ventilation&#160;and respiratory outcomes, but also in the perioperative arena as a diagnostic tool to detect phrenic nerve function. Diaphragmatic assessment has traditionally utilized fluoroscopy and nerve studies that are time-consuming, costly, and non-portable. Point-of-care ultrasound (POCUS) overcomes these barriers and can be used as a tool for non-invasive screening of diaphragm function. However, POCUS for diaphragmatic dysfunction currently suffers from several issues such as a lack of consensus guidelines, a multiplicity of protocols, and poor interoperator reliability among existing protocols, most notably with the assessment of dome of diaphragm excursion and diaphragmatic thickening. To address these issues, this manuscript reviews the available literature on diaphragmatic POCUS and identifies an image acquisition technique that is both simple to perform and has high interoperator reliability. This technique first describes a qualitative evaluation of diaphragm excursion, followed by a quantitative assessment of the excursion of the zone of apposition. The technique is described stepwise along with all the following: patient positioning, transducer selection, probe placement, image optimization, and interpretation.

Point-of-care ultrasound (POCUS) is an essential technique for screening for diaphragmatic dysfunction due to its portability, non-invasiveness, and real-time imaging capabilities. Although current diaphragmatic POCUS protocols exist, they suffer from poor interoperator reliability and lack consensus guidelines. Here we describe a technique that is reproducible and simple to perform.

Diaphragm dysfunction is a widely recognized concern across numerous medical specialties and clinical settings. Timely and accurate assessment of the ...

Medicine

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

Source: Martin, M. et al. <a href="https://app.jove.com/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.
1. ...

Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

Summary

Explore More Videos

Chapters in this video

Assessment of Motor Balance and Coordination in Mice using the Balance Beam

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

Measuring Neuromuscular Junction Functionality

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea

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

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

Diaphragmatic Ultrasound in Adults: Image Acquisition and Interpretation

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