This work was made possible thanks to financial support from the Fondation pour la Recherche M&eacute;dicale (FRM), the Milton Safenowitz PostDoctoral Fellowship for ALS Research (ALS Association), NIH NINDS Grants NS05462 and NS034382, and ANR Grant HyperMND.

1. Step One
Pre-anesthetic medication: 10-15 min before the induction of anesthesia, inject atropine (0.20 mg/kg) and methylprenidsolone (0.05 mg) sub-cutaneously to prevent salivation and oedema, respectively.
2. Step Two
Induction of anesthesia: inject pentobarbital sodium (70 mg/kg) or a mixture of ketamine/xylazine (100 mg/kg and 10 mg/kg, respectively) intra-peritoneally. Let the mouse go under until no toe pinch reflex can be obtained. If the anesthesia seems too light, supplement with 1/4 of the dose.
3. Step Three
*Note: this surgery is a terminal procedure.
When a surgical plane of anesthesia has been reached, transfer the mouse on a raised warm blanket in a prone position.
<ol>
	<li>Cover the snout of the mouse with a mask delivering pure O2 at a flow around 100 ml/min.</li>
	<li>Using a hair clipper, shave the dorsal region from the nape of the neck to the base of the tail.</li>
	<li>Shave also the right hindlimb.</li>
	<li>Flip mouse to supine position but take care to leave the oxygen mask in place.</li>
</ol>
4. Step Four
Secure the mouse in place with sutures looped around the limbs and secured at the four corners of the work surface.
5. Step Five
Insert a temperature probe to monitor the mouse core temperature. Adjust heating blanket/lamp power to maintain core temperature between 36 &deg;C and 38 &deg;C.
6. Tracheotomy and Artificial Ventilation
<ol>
	<li>Using blunt scissors, make a cut over the trachea and pull the skin away on both sides.</li>
	<li>Using blunt forceps, tear apart the salivary gland so as to expose two thin muscles (Sternohyoid) covering the trachea.</li>
	<li>Using blunt forceps, separate the two muscles along their medial separation to reveal the trachea.</li>
	<li>Using a Dumont 7 forceps, slide two length of 4.0 silk suture under the trachea.</li>
	<li>Make a transversal cut in the trachea in between two cartilaginous rings but take care not to completely section the trachea.</li>
	<li>Insert the tracheal tube down the trachea, then secure it on both sides of the insertion opening by tying the sutures over it. The tracheal tube is connected to a mouse ventilator (SAR-830/AP, CWE Inc.) and a capnograph (&mu;capstar, CWE Inc.). The ventilator is connected, through a compliance bag, to a source of pure O2. Adjust the parameters of the ventilator (breathing rate 100-150 bpm, end-tidal volume 170-310 &mu;l) so that the mouse is not fighting against the artificial ventilation and the end-tidal pCO2 is stable between 4 and 5%.</li>
</ol>
7. Placement of the Intravenous Lines
<ol>
	<li>Using blunt dissection techniques, expose the jugular vein on one side of the neck. At this level, the jugular vein splits into two major trunks, the anterior and posterior facial veins.</li>
	<li>Do the following steps twice, one set for each of these trunks:
	<ol>
		<li>Using Dumont 4 or 5 forceps, carefully separate the vein from its surrounding conjunctive tissue.</li>
		<li>Using Dumont 7 forceps, slide two length of 6.0 silk sutures under the vein. separate them as far as possible along the length of the vein.</li>
		<li>Place a small vessel clip on the proximal side of the vein (the side closest to the heart), and tie off the distal side of the vein.</li>
		<li>Using very fine iris scissors, make a small transverse incision in the vein, taking great care not to section the vein completely.</li>
		<li>Insert a pre-filled 1Fr catheter (premicath, Vygon) in the opening, up to the vessel clip.</li>
		<li>Holding the vein and the catheter together in a Dumont 4 forceps, carefully remove the vessel clip, then push the catheter a few more millimeters into the vein.</li>
		<li>Secure the catheter by tying it with both sutures on both sides of the insertion.</li>
	</ol>
	</li>
	<li>One of the catheters is connected to a syringe or a syringe pump to inject supplemental doses of anesthetics whenever necessary (usually every 10-30 min). The IV dose is either 6 mg/kg of pentobarbital sodium or 1250+40 &mu;g/kg/min of ketamine/xylazine12. The other catheter is connected to a syringe pump for a slow intravenous infusion (50 &mu;l/hr) of a 4 % glucose solution containing NaHCO3 (1 %) and plasmion (14 %).</li>
</ol>
8. Close the Neck Skin with Needle and Suture, and Return the Mouse to Prone Position
9. Dissection of the Hind-limb Muscle and Nerves
<ol>
	<li>Using scissors, make an incision from the top of the thigh to the Achilles tendon. Separate the skin from the underlying muscles, taking care not to damage blood vessels. Cauterize as needed.</li>
	<li>Identify the anterior edge of the Biceps Femoris, which appears as a white line running down the thigh. Using scissors, carefully open along the line from the knee all the way to the hip bone. Separate the muscles. The sciatic nerve is now visible under the Biceps Femoris.</li>
	<li>Carefully dissect the Biceps Femoris. Cauterize/ligature as needed to prevent bleeding. The Biceps Femoris can be totally removed or simply reclined to expose the sciatic nerve and the Triceps Surae muscles.</li>
	<li>Dissect out the Sural nerve.</li>
	<li>Using 8/0 silk thread, tie up the distal part of the Common Peroneal nerve and cut distal to the knot. Dissect the nerve all the way up, as close to the hip as possible.</li>
	<li>Identify the Tibial nerve in between the Common Peroneal and Sural nerves. Among the different branches of the Tibial nerve, identify the branches innervating the Triceps Surae from the branches that go deeper (henceforth referred to as Tibial nerve).</li>
	<li>Using 8/0 silk thread, tie together all the branches of the Tibial nerve while keeping intact the branches innervating the Triceps Surae. Cut the the Tibial nerve distally to the knot and dissect the nerve up as far as possible.</li>
	<li>Cover the whole hindlimb with a gauze imbibed with saline to prevent dessication while proceeding with the next step.</li>
</ol>
10. Laminectomy
<ol>
	<li>Using scissors, make an incision along the backbone. Separate the skin from the underlying muscles.</li>
	<li>Make two incisions on each side of the vertebrae to separate the subcutaneous muscles. Then cut each tendon of the muscles that attach on the side of the vertebrae on each side.</li>
	<li>Using blunt forceps and a fine curette, remove the remainder of the muscle on the dorsal side of the vertebrae around the spinous processes to clearly identify each individual vertebra.</li>
	<li>Identify vertebrae T13 and L1. T13 is the last vertebra to have ribs attached. The vertebral column will be immobilized using the Cunningham Spinal Cord vertebral clamps (Stoelting Co.). Place the spinal clamps on each side of T13 and L1. Be careful not to compress the spinal cord, but put a little bit of tension in the longitudinal axis. Check that the spinal column is well secured by pressing down gently with forceps.</li>
	<li>Using fine rongeurs, remove the spinal processes then the laminae over T13 and L1, thereby exposing the spinal cord.</li>
	<li>Cover the exposed spinal cord with small bits of cotton or spongel imbibed with saline to prevent desiccation.</li>
	<li>Place a custom made plastic bath on top of the back, surrounding the spinal cord. Secure the bath in place using 4/0 silk thread. Ensure that the bath is waterproof by sealing it with Kwik-Cast sealant (WPI).</li>
	<li>Once the Kwik-Cast has dried, remove the cotton covering the spinal cord, and fill the bath with mineral oil.</li>
	<li>Using Dumont 5 forceps and very fine iris scissors, gently pull on the dura mater surrounding the spinal cord, and open it as far as possible in both direction. Fold the dura on each side of the spinal cord.</li>
	<li>Lower the raised platform on which the mouse is lying so as to keep it suspended by the vertebral clamp.</li>
	<li>Use another vertebral clamp to clamp a spinous process of the sacral region to support the back region of the animal.</li>
	<li>A third clamp is used to immobilize the right hindlimb and ankle, flexed at a 90&deg; angle at the knee.</li>
</ol>
11. Achilles Tendon Dissection
<ol>
	<li>With the limb bent at 90&deg; at the knee and at the ankle, remove the gauze covering the hindlimb area, and dissect the Achilles tendon free of surrounding tissue. Dissect as much as possible the Triceps Surae from surrounding tissue as well.</li>
	<li>Cut the tendon of the Plantaris muscle close to the calcaneus, then cut it again to remove the tendon completely.</li>
	<li>Using a threaded needle, attach 6/0 silk thread through the Achilles tendon and make a triple knot around the tendon.</li>
	<li>Place the force transducer close to the knot, cut the distal part of the tendon, and attach it to the force transducer using the silk thread.</li>
	<li>Insert two lengths of stainless steel wire under the fascia of the Triceps Surae muscles. These wires are connected to an extracellular AC amplifier for EMG recording.</li>
	<li>Place the Common Peroneal nerve and the Tibial nerve on two bipolar hook electrodes.</li>
	<li>Place the Triceps Surae nerve on the cathode of a hook electrode, with the anode touching a nearby muscle.</li>
	<li>Connect all stimulating electrodes to an isolation unit.</li>
	<li>Place a ball electrode on the dorsal surface of the spinal cord, connected to an extracellular AC amplifier to record cord dorsum potentials. Place an Ag/AgCl reference electrode in contact with a back muscle.</li>
</ol>
12. Step Twelve
Stimulate the Triceps Surae nerve using a 50 &mu;sec square pulse of increasing intensity at a low frequency (&lt;1 Hz) until maximum twitch amplitude is observed. Slowly move the force transducer to stretch the muscle while monitoring the amplitude of the twitch response until the twitch amplitude reaches a maximum.
13. Intracellular Recordings of Motoneurons
From this point on, standard electrophysiological techniques are used to prepare an intracellular electrode, penetrate a neuron in the spinal cord and identify it as a motoneuron.
<ol>
	<li>Pull a glass micropipette to a ~1 &mu;m tip using a pipette puller (P-97 Micropipette Puller, Sutter Instruments). Fill the electrode with a KCl 3M solution (resistance of the electrode 10-20 M&Omega;).</li>
	<li>Using a micropositioner, drive the micropipette, connected to an intracellular amplifier (Axoclamp 2B, Axon Instruments) into the spinal cord. Monitor the local field potentials elicited by the stimulation of each nerve to locate the motor pool of the Triceps Surae.</li>
	<li>Carefully approach putative motoneurons while monitoring the resistance of the microelectrode. When pushing against a membrane, the resistance increases. Penetration can sometime be facilitated by using the "buzz" function of the intracellular amplifier.</li>
</ol>
14. Euthanasia Procedure
At the end of the experiment, the animal is euthanized by an overdose of pentobarbital (210 mg/kg IV), followed by decapitation.

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No conflicts of interest declared.

The preparation described here is the first that allows, in the adult mouse, simultaneous intracellular recording of a lumbar motoneuron and the measurement of the force produced by the muscle fibers innervated by its axon.
Because of the small size of the animal, the surgical skills required for this preparation can be challenging to acquire. However, once those skills are mastered, the whole surgery can be performed in three hours, and the animals can survive for up to 7 more hours after th...

A correction was made to <a href="http://jove.unicartagenaproxy.elogim.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>. There was an error in the name of one author, Marin Manuel. The author's name has been corrected to:
Marin Manuel 
 
instead of: 
 
Manuel Marin

A correction to author list has been made for article <a href="http://jove.unicartagenaproxy.elogim.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>.

Erratum: Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In Vivo

<table><tbody><tr><td>Atropine sulfate</td><td>Aguettant</td><td></td><td></td></tr><tr><td>Methylprenidsolone</td><td>Pfizer</td><td>Solu-Medrol</td><td></td></tr><tr><td>Sodium pentobarbitone</td><td>Sanofi-Aventis</td><td>Pentobarbital</td><td></td></tr><tr><td>Ketamine</td><td></td><td></td><td></td></tr><tr><td>Xylazine</td><td></td><td></td><td></td></tr><tr><td>Glucose</td><td></td><td></td><td></td></tr><tr><td>Plasma expander</td><td>Roger Bellon</td><td>Plasmagel</td><td></td></tr><tr><td>Blunt scissors</td><td>FST</td><td>14079-10</td><td></td></tr><tr><td>Blunt fine scissors</td><td>FST</td><td>15025-10</td><td></td></tr><tr><td>Vannas Spring Scissors</td><td>FST</td><td>15002-08</td><td></td></tr><tr><td>Fine forceps serrated</td><td>FST</td><td>11370-32</td><td></td></tr><tr><td>Fine forceps serrated</td><td>FST</td><td>11370-31</td><td></td></tr><tr><td>Cunningham Spinal Adaptor</td><td>Stoelting Co.</td><td></td><td></td></tr><tr><td>Kwik-Cast sealant</td><td>WPI</td><td>&lt;strong&gt;#&lt;/strong&gt;KWIK-CAST</td><td></td></tr><tr><td>Ventilator</td><td>CWE Inc</td><td>SAR-830/AP</td><td></td></tr><tr><td>Capnograph</td><td>CWE Inc</td><td>&amp;#956;capstar</td><td></td></tr><tr><td>Heating blanket</td><td>Harvard Apparatus</td><td>507221F</td><td></td></tr><tr><td>Intracellular amplifier</td><td>Axon Instruments</td><td>Axoclamp 2B</td><td></td></tr><tr><td>Pipette puller</td><td>Sutter Instruments</td><td>P-97</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P9333-500G</td><td></td></tr></tbody></table>

simultaneous intracellular recording of a lumbar motoneuron and the force produced by its motor unit in the adult mouse in vivo

The spinal motoneuron has long been a good model system for studying neural function because it is a neuron of the central nervous system with the unique properties of (1) having readily identifiable targets (the muscle fibers) and therefore having a very well-known function (to control muscle contraction); (2) being the convergent target of many spinal and descending networks, hence the name of "final common pathway"; and (3) having a large soma which makes it possible to penetrate them with sharp intracellular electrodes. Furthermore, when studied in vivo, it is possible to record simultaneously the electrical activity of the motoneurons and the force developed by their muscle targets. Performing intracellular recordings of motoneurons in vivo therefore put the experimentalist in the unique position of being able to study, at the same time, all the compartments of the "motor unit" (the name given to the motoneuron, its axon, and the muscle fibers it innervates1): the inputs impinging on the motoneuron, the electrophysiological properties of the motoneuron, and the impact of these properties on the physiological function of the motoneurons, i.e. the force produced by its motor unit. However, this approach is very challenging because the preparation cannot be paralyzed and thus the mechanical stability for the intracellular recording is reduced. Thus, this kind of experiments has only been achieved in cats and in rats. However, the study of spinal motor systems could make a formidable leap if it was possible to perform similar experiments in normal and genetically modified mice.
For technical reasons, the study of the spinal networks in mice has mostly been limited to neonatal in vitro preparations, where the motoneurons and the spinal networks are immature, the motoneurons are separated from their targets, and when studied in slices, the motoneurons are separated from most of their inputs. Until recently, only a few groups had managed to perform intracellular recordings of motoneurons in vivo2-4 , including our team who published a new preparation which allowed us to obtain very stable recordings of motoneurons in vivo in adult mice5,6. However, these recordings were obtained in paralyzed animals, i.e. without the possibility to record the force output of these motoneurons. Here we present an extension of this original preparation in which we were able to obtain simultaneous recordings of the electrophysiological properties of the motoneurons and of the force developed by their motor unit. This is an important achievement, as it allows us to identify the different types of motoneurons based on their force profile, and thereby revealing their function. Coupled with genetic models disturbing spinal segmental circuitry7-9, or reproducting human disease10,11, we expect this technique to be an essential tool for the study of spinal motor system.

Figure 1 shows how to identify a motoneuron from the Triceps Surae group after penetration. At low stimulation intensity, only a monosynaptic EPSP can be observed (Figure 1A). At higher intensity, the EPSP might be large enough to trigger an "orthodromic" spike (Figure 1B). At even higher stimulation intensity, an all-or-none antidromic spike appears, with a shorter latency than the monosynaptic EPSP (Figure 1C). If enough current is injected through the...

Watch this Scientific Journal Video about Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo at JoVE.com

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

This new method permits the simultaneous intracellular recording of a single adult mouse motoneuron and the measurement of the force produced by its muscle fibers. The combined investigation of the electrical and mechanical properties of motor units in normal and genetically modified animals is a breakthrough for the study of the neuromuscular system.

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

The  spinal motoneuron has long been a good model system for studying neural  function because it is a neuron of the central nervous system with the ...

simultaneous-intracellular-recording-lumbar-motoneuron-force-produced

Universidad de Cartagena UDEC

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14.6K Views.  Northwestern University Feinberg School of Medicine. This new method permits the simultaneous intracellular recording of a single adult mouse motoneuron and the measurement of the force produced by its muscle fibers. The combined investigation of the electrical and mechanical properties of motor units in normal and genetically modified animals is a breakthrough for the study of the neuromuscular system.

Video: Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

Electrophysiological Motor Unit Number Estimation (MUNE) Measuring Compound Muscle Action Potential (CMAP) in Mouse Hindlimb Muscles

Compound muscle action potential (CMAP) and motor unit number estimation (MUNE) are electrophysiological techniques that can be used to monitor the functional status of a motor unit pool in vivo. These measures can provide insight into the normal development and degeneration of the neuromuscular system. These measures have clear translational potential because they are routinely applied in diagnostic and clinical human studies. We present electrophysiological techniques similar to those employed in humans to allow recordings of mouse sciatic nerve function. The CMAP response represents the electrophysiological output from a muscle or group of muscles following supramaximal stimulation of a peripheral nerve. MUNE is an electrophysiological technique that is based on modifications of the CMAP response. MUNE is a calculated value that represents the estimated number of motor neurons or axons (motor control input) supplying the muscle or group of muscles being tested. We present methods for recording CMAP responses from the proximal leg muscles using surface recording electrodes following the stimulation of the sciatic nerve in mice. An incremental MUNE technique is described using submaximal stimuli to determine the average single motor unit potential (SMUP) size. MUNE is calculated by dividing the CMAP amplitude (peak-to-peak) by the SMUP amplitude (peak-to-peak). These electrophysiological techniques allow repeated measures in both neonatal and adult mice in such a manner that facilitates rapid analysis and data collection while reducing the number of animals required for experimental testing. Furthermore, these measures are similar to those recorded in human studies allowing more direct comparisons.

Electrophysiological Motor Unit Number Estimation MUNE Measuring Compound Muscle Action Potential CMAP in Mouse Hindlimb Muscles

We present refined protocols that allow in vivo monitoring of motor unit function in the mouse. Techniques to measure compound muscle action potential (CMAP) and motor unit number estimation (MUNE) in the mouse hind limb muscles innervated by the sciatic nerve are described.

Compound muscle action potential (CMAP) and motor unit number estimation (MUNE) are electrophysiological techniques that can be used to monitor the ...

Behavior

The Preparation of Oblique Spinal Cord Slices for Ventral Root Stimulation

Electrophysiological recordings from spinal cord slices have proven to be a valuable technique to investigate a wide range of questions, from cellular to network properties. We show how to prepare viable oblique slices of the spinal cord of young mice (P2 - P11). In this preparation, the motoneurons retain&#160;their axons coming out from the ventral roots of the&#160;spinal cord. Stimulation of these axons elicits back-propagating action potentials invading the motoneuron&#160;somas and exciting the motoneuron&#160;collaterals within the spinal cord. Recording of antidromic action potentials is an immediate, definitive and elegant way to characterize motoneuron identity, which surpasses&#160;other identification methods. Furthermore, stimulating the motoneuron collaterals is a simple and reliable way to excite the collateral targets of the motoneurons within the spinal cord, such as other motoneurons or Renshaw cells. In this protocol, we present antidromic recordings from the motoneuron somas as well as Renshaw cell excitation, resulting from ventral root stimulation.

We show how to prepare oblique slices of the spinal cord in young mice. This preparation allows for the stimulation of the ventral roots.

Electrophysiological recordings from spinal cord slices have proven to be a valuable technique to investigate a wide range of questions, from cellular to ...

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings provide a sensitive approach to measure nerve conduction in humans and rodent models. To broaden the technical possibilities for electromyography in mice, the measurement of compound muscle action potentials (CMAPs) from the brachial plexus nerve in the forelimb using needle electrodes is described here. CMAP recordings after stimulating the sciatic nerve in hindlimbs have been previously described. The newly introduced method here allows for the evaluation of the nerve conductivity at an additional site, and thus provides a more profound overview of the neuromuscular functionality. The technique provides information on both the relative number of functional axons and the myelination level. Thereby, this method can be applied to assess both axonal diseases as well as demyelinating conditions. This minimally invasive method does not require extraction of the nerve and therefore it is suitable for repeated measurements for longitudinal follow-up in the same animal. Similar recordings are performed in clinical setups to emphasize the translational relevance of the method.

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

The measurement of nerve conduction is a useful tool to assess mouse models of neurodegeneration but it is frequently only applied to stimulate the sciatic nerve in hindlimbs. Here, we describe a technique to measure compound muscle action potential (CMAP) in vivo in the mouse forelimb muscles innervated by the brachial plexus.

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings ...

In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological characteristics in the intact spinal network and holds significant advantages relative to classical in vitro or extracellular recording techniques. An advantage of in vivo intracellular recordings is that this method can be performed on adult animals with a fully mature nervous system, and therefore many observed physiological mechanisms can be translated to practical applications. In this methodological paper, we describe this procedure combined with externally applied constant current stimulation, which mimics polarization processes occurring within spinal neuronal networks. Trans-spinal direct current stimulation (tsDCS) is an innovative method increasingly used as a neuromodulatory intervention in rehabilitation after various neurological injuries as well as in sports. The influence of tsDCS on the nervous system remains poorly understood and the physiological mechanisms behind its actions are largely unknown. The application of the tsDCS simultaneously with intracellular recordings enables us to directly observe changes of motoneuron membrane properties and characteristics of rhythmic firing in response to the polarization of the spinal neuronal network, which is crucial for the understanding of tsDCS actions. Moreover, when the presented protocol includes the identification of the motoneuron with respect to an innervated muscle and its function (flexor versus extensor) as well as the physiological type (fast versus slow) it provides an opportunity to selectively investigate the influence of tsDCS on identified components of spinal circuitry, which seem to be differently affected by polarization. The presented procedure focuses on surgical preparation for intracellular recordings and stimulation with an emphasis on the steps which are necessary to achieve preparation stability and reproducibility of results. The details of the methodology of the anodal or cathodal tsDCS application are discussed while paying attention to practical and safety issues.

This protocol describes in vivo intracellular recording of rat lumbar motoneurons with simultaneous trans-spinal direct current stimulation. The method enables us to measure membrane properties and to record rhythmic firing of motoneurons before, during and after anodal or cathodal polarization of the spinal cord.

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological ...

Functional Isolation of Single Motor Units of Rat Medial Gastrocnemius Muscle

This work outlines functional isolation of motor units (MUs), a standard electrophysiological method for determining characteristics of motor units in hindlimb muscles (such as the medial gastrocnemius, soleus, or plantaris muscle) in experimental rats. A crucial element of the method is the application of electrical stimuli delivered to a motor axon isolated from the ventral root. The stimuli may be delivered at constant or variable inter-pulse intervals. This method is suitable for experiments on animals at varying stages of maturity (young, adult or old). Moreover, this protocol can be used in experiments studying variability and plasticity of motor units evoked by a large spectrum of interventions. The results of these experiments may both augment basic knowledge in muscle physiology and be translated into practical applications. This procedure focuses on the surgical preparation for the recording and stimulation of MUs, with an emphasis on the necessary steps to achieve preparation stability and reproducibility of results.

This method allows the recording of the force of twitch and tetanic contractions and action potentials in three types of motor units in the rat medial gastrocnemius muscle. The functional isolation of a single motor unit is induced by electrical stimulation of the axon.

This work outlines functional isolation of motor units (MUs), a standard electrophysiological method for determining characteristics of motor units in ...

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an increasingly detailed view of the circuits underpinning spinal pain processing. However, the effects of these connections for broader network activity in the DH remain less well understood because most studies focus on the activity of single neurons and small microcircuits. Alternatively, the use of microelectrode arrays (MEAs), which can monitor electrical activity across many cells, provides high spatial and temporal resolution of neural activity. Here, the use of MEAs with mouse spinal cord slices to study DH activity induced by chemically stimulating DH circuits with 4-aminopyridine (4-AP) is described. The resulting rhythmic activity is restricted to the superficial DH, stable over time, blocked by tetrodotoxin, and can be investigated in different slice orientations. Together, this preparation provides a platform to investigate DH circuit activity in tissue from na&#239;ve animals, animal models of chronic pain, and mice with genetically altered nociceptive function. Furthermore, MEA recordings in 4-AP-stimulated spinal cord slices can be used as a rapid screening tool to assess the capacity of novel antinociceptive compounds to disrupt activity in the spinal cord DH.

The combined use of microelectrode array technology and 4-aminopyridine-induced chemical stimulation for investigating network-level nociceptive activity in the spinal cord dorsal horn is outlined.

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an ...

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in the brain sense various biological conditions in the periphery and transmit the signals to neurons. Microglia, immune cells in the brain, are involved in synaptic development and plasticity. Therefore, the contribution of microglia to neural circuit construction in response to the internal state of the body should be tested critically by intravital imaging of the relationship between microglial dynamics and neuronal activity.
Here, we describe a technique for the simultaneous imaging of microglial dynamics and neuronal activity in awake mice. Adeno-associated virus encoding R-CaMP, a gene-encoded calcium indicator of red fluorescence protein, was injected into layer 2/3 of the primary visual cortex in CX3CR1-EGFP transgenic mice expressing EGFP in microglia. After viral injection, a cranial window was installed onto the brain surface of the injected region. In vivo two-photon imaging in awake mice 4 weeks after the surgery demonstrated that neural activity and microglial dynamics could be recorded simultaneously at the sub-second temporal resolution. This technique can uncover the coordination between microglial dynamics and neuronal activity, with the former responding to peripheral immunological states and the latter encoding the internal brain states.

Here, we describe a protocol combining adeno-associated virus injection with cranial window implantation for simultaneous imaging of microglial dynamics and neuronal activity in awake mice.

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in ...

Fabrication and Surgical Implantation of Electromyographic Electrodes in Mice

Recording Forelimb Muscle Activity in Head&#45;Fixed Mice with Chronically Implanted EMG Electrodes

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with unprecedented precision in head-fixed mice performing a variety of tasks. The small size of the mouse makes the measurement of motor output difficult, as the traditional method of electromyographic (EMG) recording of muscle activity was designed for larger animals like cats and primates. Pending commercially available EMG electrodes for mice, the current gold-standard method for recording muscle activity in mice is to make electrode sets in-house. This article describes a refinement of established procedures for hand fabrication of an electrode set, implantation of electrodes in the same surgery as headplate implantation, fixation of a connector on the headplate, and post-operative recovery care. Following recovery, millisecond-resolution EMG recordings can be obtained during head-fixed behavior for several weeks without noticeable changes in signal quality. These recordings enable precise measurement of forelimb muscle activity alongside in vivo neural recording and/or perturbation to probe mechanisms of motor control in mice.

Author Spotlight: Investigating Mouse Motor Cortex Interactions from Muscle Activity to Neural Dynamics

This protocol describes the hand fabrication and surgical implantation of electromyographic (EMG) electrodes in the forelimb muscles of mice to record muscle activity during head-fixed behavior experiments.

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with ...

Trans-Spinal Direct Current Stimulation of a Rat&#39;s Spinal Motoneuron

Source: B&#261;czyk, M., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/61439">In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation.</a> J. Vis. Exp. (2020).
This video describes the in vivo intracellular recording of rat motoneurons under trans-spinal direct current stimulation. The protocol describes how to measure membrane properties and record the rhythmic firing of motoneurons before, during, and after anodal or cathodal polarization of the spinal cord.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
NOTE: ...

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

Measuring Compound Muscle Action Potentials in Mouse Forelimb Muscles In Vivo

Source: Pollari, E. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/57741">In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration.</a>&#160;J. Vis. Exp. (2018).
This video demonstrates an in vivo technique for measuring compound muscle action potential (CMAP) in mice. It outlines the steps for setting up electrodes, stimulating forelimb neurons that innervate muscle fibers, and recording muscle responses. The technique assesses neuronal demyelination and functional neuron loss.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Animal ...

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

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Chapters in this video

Electrophysiological Motor Unit Number Estimation (MUNE) Measuring Compound Muscle Action Potential (CMAP) in Mouse Hindlimb Muscles

The Preparation of Oblique Spinal Cord Slices for Ventral Root Stimulation

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

Functional Isolation of Single Motor Units of Rat Medial Gastrocnemius Muscle

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Recording Forelimb Muscle Activity in Head-Fixed Mice with Chronically Implanted EMG Electrodes

Trans-Spinal Direct Current Stimulation of a Rat's Spinal Motoneuron

Measuring Compound Muscle Action Potentials in Mouse Forelimb Muscles In Vivo