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 o...

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 border="0" cellpadding="0" cellspacing="0" width="902">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Paraformaldehyde</td>
			<td style="width:273px;">Fisher</td>
			<td style="width:131px;">T353-500</td>
			<td style="width:167px;">Make 10% solution first in de-ionized distilled water; make 4% with 1X PBS, adjust pH to 7.4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">1X Phosphate Buffered Saline, pH 7.4</td>
			<td style="width:273px;">Invitrogen</td>
			<td style="width:131px;">10010049</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">2% Bovine serum albumin (2% BSA)</td>
			<td style="width:273px;">Sigma-Aldrich</td>
			<td style="width:131px;">A3059-100g</td>
			<td>Dissolve 2g BSA into 100mL of 1X PBS</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">0.2% Triton X100 in 2% BSA/PBS (Blocking Buffer)</td>
			<td style="width:273px;">Sigma-Aldrich</td>
			<td style="width:131px;">T9284-100mL</td>
			<td>Dissolve 0.2ml/100mL 2% BSA/PBS</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">0.1M Glycine</td>
			<td style="width:273px;">Sigma-Aldrich</td>
			<td style="width:131px;">G-7126</td>
			<td>Add 0.185g to 25mL of 2% BSA/PBS</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">&#945;-bungarotoxin</td>
			<td style="width:273px;">Invitrogen</td>
			<td style="width:131px;">T1175</td>
			<td>Concentration 1:400</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">SMI-312&#160;</td>
			<td style="width:273px;">Sternberger Monoclonals</td>
			<td style="width:131px;">SMI312</td>
			<td>Concentration 1:1,000</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">SV2</td>
			<td style="width:273px;">Developmental Studies Hybridoma Bank</td>
			<td style="width:131px;">SV2-Supernatant</td>
			<td>Concentration 1:10</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">FITC goat anti-mouse IgG1</td>
			<td style="width:273px;">Roche</td>
			<td style="width:131px;">3117731001</td>
			<td>Concentration 1:100</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Silicone rubber</td>
			<td style="width:273px;">Sylgard, Dow Corning</td>
			<td style="width:131px;">Part # 184</td>
			<td>Follow instructions that come with kit: can use multiple sized culture dish (30mm, 60mm, 100mm) depending on needs</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Vectashield fluorescent mounting medium</td>
			<td style="width:273px;">Vector laboratories</td>
			<td style="width:131px;">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 height="21">
			<td height="21" style="height:21px;width:332px;">Small Spring Scissors</td>
			<td style="width:273px;">Fine Science Tools</td>
			<td style="width:131px;">15002-08</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Dissection forceps</td>
			<td style="width:273px;">Fine Science Tools</td>
			<td style="width:131px;">11295-51</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Software for CMAP recordings</td>
			<td style="width:273px;">Scope 3.5.6; ADI</td>
			<td style="width:131px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Disk surface electrodes</td>
			<td style="width:273px;">Natus neurology</td>
			<td style="width:131px;">019-409000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Subdermal needle electrodes</td>
			<td style="width:273px;">Natus neurology</td>
			<td style="width:131px;">019-453100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Conductive gel</td>
			<td style="width:273px;">Aquasonic&#160;</td>
			<td style="width:131px;">122-73720</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:332px;">Stimulator/recording system for CMAP recordings</td>
			<td style="width:273px;">ADI Powerlab 8SP stimulator&#160;</td>
			<td style="width:131px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:332px;">Amplifier for CMAP recordings</td>
			<td style="width:273px;">BioAMP</td>
			<td style="width:131px;"></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<...

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

Research

JoVE Journal

Neuroscience

17.4K 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 ...

Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, direction5-7, ocular dominance8,9 and binocular disparity9. Orientation selectivity is the most widely studied feature and a continuous map with a quasi-periodic layout for preferred orientation is present across the entire primary visual cortex10,11. Integrating the synaptic, cellular and network contributions that lead to stimulus selective responses in these functional maps requires the hybridization of imaging techniques that span sub-micron to millimeter spatial scales. With conventional intrinsic signal optical imaging, the overall layout of functional maps across the entire surface of the visual cortex can be determined12. The development of in vivo two-photon microscopy using calcium sensitive dyes enables one to determine the synaptic input arriving at individual dendritic spines13 or record activity simultaneously from hundreds of individual neuronal cell bodies6,14. Consequently, combining intrinsic signal imaging with the sub-micron spatial resolution of two-photon microscopy offers the possibility of determining exactly which dendritic segments and cells contribute to the micro-domain of any functional map in the neocortex. Here we demonstrate a high-yield method for rapidly obtaining a cortical orientation map and targeting a specific micro-domain in this functional map for labeling neurons with fluorescent dyes in a non-rodent mammal. With the same microscope used for two-photon imaging, we first generate an orientation map using intrinsic signal optical imaging. Then we show how to target a micro-domain of interest using a micropipette loaded with dye to either label a population of neuronal cell bodies or label a single neuron such that dendrites, spines and axons are visible in vivo. Our refinements over previous methods facilitate an examination of neuronal structure-function relationships with sub-cellular resolution in the framework of neocortical functional architectures.

A method is described for labeling neurons with fluorescent dyes in predetermined functional micro-domains of the neocortex. First, intrinsic signal optical imaging is used to obtain a functional map. Then two-photon microscopy is used to label and image neurons within a micro-domain of the map.

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, ...

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 ...

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 ...

Conversion of Human Induced Pluripotent Stem Cells (iPSCs) into Functional Spinal and Cranial Motor Neurons Using PiggyBac Vectors

We describe here a method to obtain functional spinal and cranial motor neurons from human induced pluripotent stem cells (iPSCs). Direct conversion into motor neuron is obtained by ectopic expression of alternative modules of transcription factors, namely Ngn2, Isl1 and Lhx3 (NIL) or Ngn2, Isl1 and Phox2a (NIP). NIL and NIP specify, respectively, spinal and cranial motor neuron identity. Our protocol starts with the generation of modified iPSC lines in which NIL or NIP are stably integrated in the genome via a piggyBac transposon vector. Expression of the transgenes is then induced by doxycycline and leads, in 5 days, to the conversion of iPSCs into MN progenitors. Subsequent maturation, for 7 days, leads to homogeneous populations of spinal or cranial MNs. Our method holds several advantages over previous protocols: it is extremely rapid and simplified; it does not require viral infection or further MN isolation; it allows generating different MN subpopulations (spinal and cranial) with a remarkable degree of maturation, as demonstrated by the ability to fire trains of action potentials. Moreover, a large number of motor neurons can be obtained without purification from mixed populations. iPSC-derived spinal and cranial motor neurons can be used for in vitro modeling of Amyotrophic Lateral Sclerosis and other neurodegenerative diseases of the motor neuron. Homogeneous motor neuron populations might represent an important resource for cell type specific drug screenings.

Conversion of Human Induced Pluripotent Stem Cells iPSCs into Functional Spinal and Cranial Motor Neurons Using PiggyBac Vectors

This protocol allows rapid and efficient conversion of induced pluripotent stem cells into motor neurons with a spinal or cranial identity, by ectopic expression of transcription factors from inducible piggyBac vectors.

We describe here a method to obtain functional spinal and cranial motor neurons from human induced pluripotent stem cells (iPSCs). Direct conversion into ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

Axon degeneration is a shared feature in neurodegenerative disease and when nervous systems are challenged by mechanical or chemical forces. However, our understanding of the molecular mechanisms underlying axon degeneration remains limited. Injury-induced axon degeneration serves as a simple model to study how severed axons execute their own disassembly (axon death). Over recent years, an evolutionarily conserved axon death signaling cascade has been identified from flies to mammals, which is required for the separated axon to degenerate after injury. Conversely, attenuated axon death signaling results in morphological and functional preservation of severed axons and their synapses. Here, we present three simple and recently developed protocols that allow for the observation of axonal morphology, or axonal and synaptic function of severed axons that have been cut-off from the neuronal cell body, in the fruit fly Drosophila. Morphology can be observed in the wing, where a partial injury results in axon death side-by-side of uninjured control axons within the same nerve bundle. Alternatively, axonal morphology can also be observed in the brain, where the whole nerve bundle undergoes axon death triggered by antennal ablation. Functional preservation of severed axons and their synapses can be assessed by a simple optogenetic approach coupled with a post-synaptic grooming behavior. We present examples using a highwire loss-of-function mutation and by over-expressing dnmnat, both capable of delaying axon death for weeks to months. Importantly, these protocols can be used beyond injury; they facilitate the characterization of neuronal maintenance factors, axonal transport, and axonal mitochondria.

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

Here, we provide protocols to perform three simple injury-induced axon degeneration (axon death) assays in Drosophila melanogaster to evaluate the morphological and functional preservation of severed axons and their synapses.

Axon degeneration is a shared feature in neurodegenerative disease and when nervous systems are challenged by mechanical or chemical forces. However, our ...

Culturing Rat Sympathetic Neurons from Embryonic Superior Cervical Ganglia for Morphological and Proteomic Analysis

Sympathetic neurons from the embryonic rat superior cervical ganglia (SCG) have been used as an in vitro model system for peripheral neurons to study axonal growth, axonal trafficking, synaptogenesis, dendritic growth, dendritic plasticity and nerve-target interactions in co-culture systems. This protocol describes the isolation and dissociation of neurons from the superior cervical ganglia of E21 rat embryos, followed by the preparation and maintenance of pure neuronal cultures in serum-free medium. Since neurons do not adhere to uncoated plastic, neurons will be cultured on either 12 mm glass coverslips or 6-well plates coated with poly-D-lysine. Following treatment with an antimitotic agent (Ara-C, cytosine &#946;-D-arabinofuranoside), this protocol generates healthy neuronal cultures with less than 5% non-neuronal cells, which can be maintained for over a month in vitro. Although embryonic rat SCG neurons are multipolar with 5-8 dendrites in vivo; under serum-free conditions, these neurons extend only a single axon in culture and continue to be unipolar for the duration of the culture. However, these neurons can be induced to extend dendrites in the presence of basement membrane extract, bone morphogenetic proteins (BMPs), or 10% fetal calf serum. These homogenous neuronal cultures can be used for immunocytochemical staining and for biochemical studies. This paper also describes optimized protocol for immunocytochemical staining for microtubule associated protein-2 (MAP-2) in these neurons and for the preparation of neuronal extracts for mass spectrometry.

This paper describes the isolation and culturing of embryonic rat sympathetic neurons from the superior cervical ganglia. It also provides detailed protocols for immunocytochemical staining and for preparing neuronal extracts for mass spectrometric analysis.

Sympathetic neurons from the embryonic rat superior cervical ganglia (SCG) have been used as an in vitro model system for peripheral neurons to study ...