This work was supported by Charley&#39;s Fund and NIH R01NS055099.

All methods described here have been approved by the Institutional Animal Care and Use Committee of Beth Israel Deaconess Medical Center under protocol numbers (031-2019; 025-2019). Wear proper PPE equipment to handle animals and adhere to IACUC guidelines for all animal work.
1. In vivo surface EIM
<ol>
	<li>Place the animal in an anesthesia box to induce anesthesia. 
	NOTE: For rats, 1.5%-3.5% isoflurane and 2 O2 L&#183;min-1 were used, and for mice, 2% isoflurane and 1 O2 L&#183;min-1 were used.</li>
	<li>Once fully anesthetized, as indicated by the absence of response after pinching the foot of the animal, place the mouse on the bench in a prone position and use the nose cone to maintain anesthesia using 1.5% isoflurane and an oxygen flow of 1 L&#183;min-1.</li>
	<li>Place the animal&#39;s leg to be analyzed at a 45&#176; angle with the hip joint (knee extended) and secure the foot with medical tape.</li>
	<li>Use a hair clipper to trim the fur overlaying the gastrocnemius muscle.</li>
	<li>Apply a thick layer of depilatory cream over the animal&#39;s skin and let it sit for 1 min. Then, use saline-saturated gauze to remove the depilatory agent. Repeat this process up to three times until all the fur overlaying the gastrocnemius muscle is removed. 
	NOTE: Place a gauze pad soaked in saline over the skin when measurements are not being acquired to prevent skin dehydration.</li>
	<li>Connect the surface array (Figure 1) to the EIM device and let the electrodes rest on a piece of gauze soaked in saline solution.</li>
	<li>Place the surface array directly on the skin over the gastrocnemius muscle, oriented longitudinally to the muscle fibers.</li>
	<li>After checking for appropriate contact, which is indicated by all bars appearing green on the software showing the stability of the 50 kHz resistance, reactance, and phase values, acquire the EIM measurements. 
	NOTE: Curves should be checked in real time to ensure proper data acquisition.</li>
	<li>Rotate the surface array by 90&#176; and reposition it on the skin over the gastrocnemius to obtain the transverse measurements (check for green bars indicating the stability).</li>
	<li>Repeat steps 1.7, 1.8, and 1.9 to get a total of four measurements per muscle: two longitudinal and two transverse. 
	NOTE: Do not use a depilatory agent more than once (i.e., up to three applications in the same instance) every two weeks to prevent excessive skin irritation and injury. It is important to perform the measurements within about 5-10 min of removing the depilatory cream since the development of localized skin edema induced by the depilatory agent may impact the collected impedance data. Animal recovery is immediate after stopping isoflurane anesthesia and the procedure does not require analgesic treatment.</li>
</ol>
2. In vivo needle array EIM
<ol>
	<li>Anesthetize the animal and prepare the leg using the same procedure as described in steps 1.1-1.4. However, it is not necessary to use a depilatory agent when performing in vivo EIM using a needle array.</li>
	<li>Connect the needle array (Figure 2A-F) to the EIM device and let it rest in a weighing boat containing saline solution. Check for connectivity and signal stability (indicated by green bars).</li>
	<li>Disinfect the skin and needles with alcohol. Place the needle array in a longitudinal position compared to the myofibers and press it firmly into the skin until all the needles penetrate the skin and the underlying muscle up to the plastic guard on the array. Acquire data.</li>
	<li>Gently remove the array and reinsert it through the skin and into the muscle at a 90&#176; angle relative to the first measurement, in the transverse direction. Acquire data. 
	NOTE: When using needle arrays, measurements should only be acquired once in each direction to reduce the impact of the needle electrodes on the skin and muscle tissue. If bleeding occurs, gently wipe the blood away before performing the second measurement. Animal recovery is immediate after stopping isoflurane anesthesia and the procedure does not require analgesic treatment.</li>
</ol>
3. Ex vivo EIM
<ol>
	<li>Prepare the ex vivo dielectric cell (Figure 2G,H), add saline solution to the chamber, and connect the cell to the EIM device to obtain the reference values. 
	NOTE: The phase and reactance values of saline should remain constant at or near zero and the resistance values of saline should remain constant at approximately 100 &#177; 25 &#937; over the frequency range from 1 kHz to 1 MHz.</li>
	<li>Euthanize the animal according to respective IACUC guidelines.</li>
	<li>Using a pair of scissors, cut the skin near the Achilles tendon. Using tweezers, pull the skin in an upward motion to reveal the underlying muscles and fascia. Gently dissect out the biceps femoris overlaying the gastrocnemius muscle and section the sciatic nerve.</li>
	<li>Cut the Achilles tendon to free the distal end of the gastrocnemius and soleus muscles and gently pull the tendon upwards while using scissors to remove any attachments. Once all attachments are removed, use scissors to cut the rostral end of the soleus muscle and remove it.</li>
	<li>Use scissors to dissect the heads of the gastrocnemius muscle around the patella. 
	NOTE: After removal of the gastrocnemius muscle, it is important to remember the original orientation of the myofibers.</li>
	<li>Place the gastrocnemius muscle on a sheet of dental wax and section it using a razor blade and a ruler to obtain a 10 mm x 10 mm section from the center of the gastrocnemius muscle. 
	NOTE: The dielectric cell size can be customized. For rats, a 10 mm x 10 mm cell was used and for mice, a 5 mm x 5 mm cell was used.</li>
	<li>Using tweezers, gently place the gastrocnemius in the dielectric cells, making sure the fibers are oriented longitudinally (i.e., caudal, and rostral extremities should be touching the electrodes). Make sure that the muscle is fully in contact with the metal electrodes.</li>
	<li>Attach the top part of the dielectric cell and insert two monopolar needles (26 G) into the two holes. Connect the wires from the EIM device to the ex vivo cell in the following order: (1: I+, 2: V+, 3: V-, 4: I-, where I represents the current electrodes and V represents the voltage electrodes). Acquire the longitudinal measurement.</li>
	<li>Open the dielectric cell and reorient the muscle in the transverse direction by rotating it 90&#176;. Reattach the top of the dielectric cell. Acquire the transverse measurement.</li>
</ol>

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S. B. Rutkove has equity in, and serves as a consultant and scientific advisor to, Myolex, Inc., a company that designs impedance devices for clinical and research use, and the mView system used here. He is also a member of the company's Board of Directors. The company also has an option to license patented impedance technology of which S. B. Rutkove is named as an inventor. The other authors have no other relevant affiliations or financial involvement with any organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript apart from those disclosed.

This article provides the basic methods for performing EIM in rodents, both in vivo and ex vivo. To acquire reliable measurements, it is critical to perform a series of steps. First, one needs to properly identify the muscle of interest, as each muscle will have different responses to diseases, treatment, and pathology. One must be mindful that the data acquired on one muscle (e.g., gastrocnemius) will not provide the same information as on another muscle (e.g., tibialis anterior). Second, one needs to ...

Electrical impedance myography (EIM) provides a powerful method to assess muscle condition, potentially enabling the diagnosis of neuromuscular disorders, tracking of disease progression, and assessment of response to therapy1,2,3. It can be applied analogously to animal disease models and humans, allowing for relatively seamless translation from preclinical to clinical studies. EIM measurements are easily obtained using four linearly-placed electrodes, with the two outer ones applying a painless, weak electrical current across a range of frequencies (generally between 1 kHz and approximately 2 MHz), and the two inner ones recording the resulting voltages1. From these voltages, the impedance characteristics of the tissue can be obtained, including the resistance (R), a measure of how difficult it is for current to pass through the tissue, and the reactance (X) or &#34;chargeability&#34; of the tissue, a measure related to the tissue&#39;s ability to store charge (capacitance). From the reactance and resistance, the phase angle (&#952;) is calculated via the following equation: <img alt="Equation 1" src="/files/ftp_upload/63513/63513eq01.jpg" style="margin: auto;" />, providing a single summative impedance measure. Such measurements can be obtained using any multifrequency bioimpedance device. As myofibers are essentially long cylinders, muscle tissue is also highly anisotropic, with current flowing more easily along fibers than across them4,5. Thus, EIM is often performed in two directions: with the array placed along the fibers such that current runs parallel to them, and across the muscle such that the current flows perpendicular to them. Additionally, in ex vivo measurements, where a known volume of tissue is measured in an impedance measuring cell, the inherent electrical properties of the muscle (i.e. the conductivity and relative permittivity), can be derived6.
The term &#34;neuromuscular disorders&#34; defines a wide range of primary and secondary diseases that lead to structural muscle alteration and dysfunction. This includes amyotrophic lateral sclerosis and various forms of muscular dystrophy, as well as simpler changes related to aging (e.g., sarcopenia), disuse atrophy (e.g., due to prolonged bedrest or microgravity) or even injury7. While the causes are plentiful and can originate from the motor neuron, nerves, neuromuscular junctions, or the muscle itself, EIM can be used to detect early alterations in muscle due to many of these processes and to track progression or response to therapy. For example, in patients with Duchenne muscular dystrophy (DMD), EIM has been shown to detect disease progression and response to corticosteroids8. Recent work has also shown EIM to be sensitive to varying disuse states, including fractional gravity9, as would be experienced on the Moon or Mars, and the effects of aging10,11. Finally, by applying predictive and machine learning algorithms to the data set obtained with each measurement (multifrequency and directionally dependent data), it becomes possible to infer histological aspects of the tissue, including myofiber size12,13, inflammatory changes and edema14, and connective tissue and fat content15,16.
Several other noninvasive or minimally invasive methods are also used to evaluate muscle health in humans and animals, including needle electromyography17 and imaging technologies such as magnetic resonance imaging, computerized tomography, and ultrasound18,19. However, EIM demonstrates distinct benefits compared with these technologies. For example, electromyography records only the active electrical properties of the myofiber membranes and not the passive properties, and thus cannot provide a true assessment of muscle composition or structure. In a certain respect, imaging methods are more closely related to EIM, as they too provide information about the structure and composition of tissue. But in some sense, they provide too much data, requiring detailed image segmentation and expert analysis rather than just providing a quantitative output. Moreover, given their complexities, imaging techniques are also greatly impacted by the specifics of both the hardware and software being used, ideally requiring the use of identical systems so that data sets can be compared. In contrast, the fact that EIM is much simpler means that it is less impacted by these technical issues and does not require any form of image processing or expert analysis.
The following protocol demonstrates how to perform in vivo EIM in rats and mice, using both noninvasive (surface array) and minimally invasive (subdermal needle array) techniques, as well as ex vivo EIM on freshly excised muscle.

<table><tbody><tr><td>3D Printer</td><td>Formlabs Inc.</td><td>Form 2 Desktop</td><td>3D printer</td></tr><tr><td>3D Printer</td><td>Shenzhen Creality 3D Technology Co. LTD</td><td>Creality Ender 3 V2</td><td>3D printer</td></tr><tr><td>3M Micropore surgical tape</td><td>Fisher</td><td>19-027761 and 19-061655</td><td>models 1530-0 and 1530-1</td></tr><tr><td>3M TRANSPORE surgical tape</td><td>Fisher</td><td>18-999-380 and 18-999-381</td><td>models 1527-0 and 1527-1</td></tr><tr><td>Connector header vertical 10 POS 1 mm spacing</td><td>Digi-Key (Sullins connector solution)</td><td>S9214-ND (SMH100-LPSE-S10-ST-BK)</td><td>Plastic spacer 1 mm holes for the rat in vivo array displayed in Figure 2A</td></tr><tr><td>Cotton-tipped applicators</td><td>Fisher</td><td>22-363-172</td><td></td></tr><tr><td>Dental Wax</td><td>Fisher</td><td>NC9377103</td><td></td></tr><tr><td>Depilatory agent</td><td>NAIR</td><td>NA</td><td>hair remover lotion with softening baby oil</td></tr><tr><td>Dumont #7b Forceps</td><td>Fine Science Tools</td><td>No. 11270-20</td><td>Used for dissection, Style: #7b, Tip Shape: Curved, Tips: Standard, Tip Dimensions: 0.17 mm x 0.1 mm, Alloy/Material: Inox, Length: 11 cm</td></tr><tr><td>Electronic Digital Caliper</td><td>Fisher</td><td>14-648-17</td><td>Used to measure out the dimensions of the Gastrocnemius muscle</td></tr><tr><td>Epoxy adhesive dual cartridge 4 min work life</td><td>Devcon</td><td>series 14265, model 2217</td><td>Glue used in the rat in vivo array displayed in Figure 2A</td></tr><tr><td>Ex vivo dielectric impedance cell</td><td>Custom</td><td>NA</td><td>Dielectric cells were 3D printed in the Rutkove laboratory</td></tr><tr><td>Graefe Forceps</td><td>Fine Science Tools</td><td>No. 11051-10</td><td>Used for muscle to place and adjust, Length: 10 cm, Tip Shape: Curved, Tips: Serrated, Tip Width: 0.8 mm, Tip Dimensions: 0.8 mm x 0.7 mm, Alloy/Material</td></tr><tr><td>Hair clipper</td><td>Amazon</td><td>NA</td><td>Wahl professional animal BravMini+</td></tr><tr><td>Impedance Animal Device</td><td>Myolex</td><td>EIM1103</td><td>mView system - investigational electrical impedance myography device for use in animal research</td></tr><tr><td>In vivo needle arrays</td><td>Custom</td><td>NA</td><td>Custom arrays using 27 G subdermal needles from Ambu. The construction was finalized using a 3D printer in the Rutkove laboratory</td></tr><tr><td>In vivo surface array</td><td>Custom</td><td>NA</td><td>The in vivo surface array was printed and assembled in the Rutkove laboratory</td></tr><tr><td>Isoflurane</td><td>Patterson Veterinary Supplies</td><td>07-893-8441 (NDC: 46066-755-04)</td><td>Pivetal - 250 mL bottle</td></tr><tr><td>Non-woven gauze</td><td>Fisher</td><td>22-028-559</td><td>2 x 2 inch</td></tr><tr><td>Polystyrene Weighing Dishes</td><td>Fisher</td><td>S67090A</td><td>Dimensions (L x W x H): 88.9 mm x 88.9 mm x 25.4 mm</td></tr><tr><td>Razor Blades</td><td>Fisher</td><td>12-640</td><td>Used to cut muscle to right dimensions, Single-edge carbon steel blades</td></tr><tr><td>Student Fine Scissors</td><td>Fine Science Tools</td><td>No. 91460-11</td><td>Used for dissection, Tips: Sharp-Sharp, Alloy/Material: Student Stainless Steel, Serrated: No, Tip Shape: Straight, Cutting Edge: 20 mm, Length: 11.5 cm, Feature: Student Quality</td></tr><tr><td>Subdermal needles 27 G Neuroline</td><td>Ambu</td><td>745 12-50/24</td><td>Needles used in the rat in vivo array displayed in Figure 2A</td></tr><tr><td>Surgical Scissors - Sharp</td><td>Fine Science Tools</td><td>No. 14002-13</td><td>Used to cut skin, Tips: Sharp-Sharp, Alloy/Material: Stainless Steel, Serrated: No, Tip Shape: Straight, Cutting Edge: 42 mm, Length: 13 cm</td></tr><tr><td>TECA ELITE monopolar needle electrodes</td><td>Natus</td><td>902-DMG50-S</td><td>0.46 mm diameter (26 G). Blue hub</td></tr><tr><td>Teknova 0.9% saline solution</td><td>Fisher</td><td>S5815</td><td>1000 mL sterile</td></tr></tbody></table>

performing in vivo and ex vivo electrical impedance myography in rodents

Electrical impedance myography (EIM) is a convenient technique that can be used in preclinical and clinical studies to assess muscle tissue health and disease. EIM is obtained by applying a low-intensity, directionally focused, electrical current to a muscle of interest across a range of frequencies (i.e., from 1 kHz to 10 MHz) and recording the resulting voltages. From these, several standard impedance components, including the reactance, resistance, and phase, are obtained. When performing ex vivo measurements on excised muscle, the inherent passive electrical properties of the tissue, namely the conductivity and relative permittivity, can also be calculated. EIM has been used extensively in animals and humans to diagnose and track muscle alterations in a variety of diseases, in relation to simple disuse atrophy, or as a measure of therapeutic intervention. Clinically, EIM offers the potential to track disease progression over time and to assess the impact of therapeutic interventions, thus offering the opportunity to shorten the clinical trial duration and reduce sample size requirements. Because it can be performed noninvasively or minimally invasively in living animal models as well as humans, EIM offers the potential to serve as a novel translational tool enabling both preclinical and clinical development. This article provides step-by-step instructions on how to perform in vivo and ex vivo EIM measurements in mice and rats, including approaches to adapt the techniques to specific conditions, such as for use in pups or obese animals.

EIM can be obtained in many conditions, including surface in vivo arrays (Figure 1), needle in vivo arrays (Figure 2A-F), and ex vivo dielectric cells (Figure 2G,H).
EIM provides a near-instantaneous snapshot of the muscle condition based on the measured impedance values. Measurements are acquired swiftly and result in a simple ...

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Performing In Vivo and Ex Vivo Electrical Impedance Myography in Rodents

This article details how to perform in vivo (using surface and needle electrode arrays) and ex vivo (using a dielectric cell) electrical impedance myography on the rodent gastrocnemius muscle. It will demonstrate the technique in both mice and rats and detail the modifications available, (i.e., obese animals, pups).

Performing In Vivo and Ex Vivo Electrical Impedance Myography in Rodents

Electrical impedance myography (EIM) is a convenient technique that can be used in preclinical and clinical studies to assess muscle tissue health and ...

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Biology

2.8K Views.  Harvard Medical School - Beth Israel Deaconess Medical Center. This article details how to perform in vivo (using surface and needle electrode arrays) and ex vivo (using a dielectric cell) electrical impedance myography on the rodent gastrocnemius muscle. It will demonstrate the technique in both mice and rats and detail the modifications available, (i.e., obese animals, pups).

Video: Performing In Vivo and Ex Vivo Electrical Impedance Myography in Rodents

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Neuroscience

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

In Vitro Recording of Mesenteric Afferent Nerve Activity in Mouse Jejunal and Colonic Segments

Afferent nerves not only convey information concerning normal physiology, but also signal disturbed homeostasis and pathophysiological processes of the different organ systems from the periphery towards the central nervous system. As such, the increased activity or &#39;sensitization&#39; of mesenteric afferent nerves has been allocated an important role in the pathophysiology of visceral hypersensitivity and abdominal pain syndromes.
Mesenteric afferent nerve activity can be measured in vitro in an isolated intestinal segment that is mounted in a purpose-built organ bath and from which the splanchnic nerve is isolated, allowing researchers to directly assess nerve activity adjacent to the gastrointestinal segment. Activity can be recorded at baseline in standardized conditions, during distension of the segment or following the addition of pharmacological compounds delivered intraluminally or serosally. This technique allows the researcher to easily study the effect of drugs targeting the peripheral nervous system in control specimens; besides, it provides crucial information on how neuronal activity is altered during disease. It should be noted however that measuring afferent neuronal firing activity only constitutes one relay station in the complex neuronal signaling cascade, and researchers should bear in mind not to overlook neuronal activity at other levels (e.g., dorsal root ganglia, spinal cord or central nervous system) in order to fully elucidate the complex neuronal physiology in health and disease.
Commonly used applications include the study of neuronal activity in response to the administration of lipopolysaccharide, and the study of afferent nerve activity in animal models of irritable bowel syndrome. In a more translational approach, the isolated mouse intestinal segment can be exposed to colonic supernatants from IBS patients. Furthermore, a modification of this technique has been recently shown to be applicable in human colonic specimens.

In Vitro Recording of Mesenteric Afferent Nerve Activity in Mouse Jejunal and Colonic Segments

Mesenteric afferent nerves convey information from the gastrointestinal tract towards the brain regarding normal homeostasis as well as pathophysiology. Gastrointestinal afferent nerve activity can be assessed by mounting isolated intestinal segments with attached afferent nerves into an organ bath, isolating the nerve, and assessing basal as well as stimulated activity.

Afferent nerves not only convey information concerning normal physiology, but also signal disturbed homeostasis and pathophysiological processes of the ...

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and latency detect chronic axon loss and demyelination, respectively. Axonal excitability techniques &#34;by threshold tracking&#34; expand upon these measures by providing information regarding the activity of ion channels, pumps and exchangers that relate to acute function and may precede degenerative events. As such, the use of axonal excitability in animal models of neurological disorders may provide a useful in vivo measure to assess novel therapeutic interventions. Here we describe an experimental setup for multiple measures of motor axonal excitability techniques in the rat ulnar nerve.
The animals are anesthetized with isoflurane and carefully monitored to ensure constant and adequate depth of anesthesia. Body temperature, respiration rate, heart rate and saturation of oxygen in the blood are continuously monitored. Axonal excitability studies are performed using percutaneous stimulation of the ulnar nerve and recording from the hypothenar muscles of the forelimb paw. With correct electrode placement, a clear compound muscle action potential that increases in amplitude with increasing stimulus intensity is recorded. An automated program is then utilized to deliver a series of electrical pulses which generate 5 specific excitability measures in the following sequence: stimulus response behavior, strength duration time constant, threshold electrotonus, current-threshold relationship and the recovery cycle.
Data presented here indicate that these measures are repeatable and show similarity between left and right ulnar nerves when assessed on the same day. A limitation of these techniques in this setting is the effect of dose and time under anesthesia. Careful monitoring and recording of these variables should be undertaken for consideration at the time of analysis.

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Axonal excitability techniques provide a powerful tool to examine pathophysiology and biophysical changes that precede irreversible degenerative events. This manuscript demonstrates the use of these techniques on the ulnar nerve of anesthetized rats.

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and ...

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 ...

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 ...

 Fabricating Epimysial EMG Electrodes for Preclinical Rodent Models 

Development of a Low-cost Epimysial Electromyography Electrode: A Simplified Workflow for Fabrication and Testing

Electromyography (EMG) is a valuable diagnostic tool for detecting neuromuscular abnormalities. Implantable epimysial electrodes are commonly used to measure EMG signals in preclinical models. Although classical resources exist describing the principles of epimysial electrode fabrication, there is a sparsity of illustrative information translating electrode theory to practice. To remedy this, we provide an updated, easy-to-follow guide on fabricating and testing a low-cost epimysial electrode. 
Electrodes were made by folding and inserting two platinum-iridium foils into a precut silicone base to form the contact surfaces. Next, coated stainless steel wires were welded to each contact surface to form the electrode leads. Lastly, a silicone mixture was used to seal the electrode. Ex vivo testing was conducted to compare our custom-fabricated electrode to an industry standard electrode in a saline bath, where high levels of signal agreement (sine [intraclass correlation - ICC= 0.993], square [ICC = 0.995], triangle [ICC = 0.958]), and temporal-synchrony (sine [r = 0.987], square [r = 0.990], triangle [r= 0.931]) were found across all waveforms. Low levels of electrode impedance were also quantified via electrochemical impedance spectroscopy. 
An in vivo performance assessment was also conducted where the vastus lateralis muscle of a rat was surgically instrumented with the custom-fabricated electrode and signaling was acquired during uphill and downhill walking. As expected, peak EMG activity was significantly lower during downhill walking (0.008 &plusmn; 0.005 mV) than uphill (0.031 &plusmn; 0.180 mV, p = 0.005), supporting the validity of the device. The reliability and biocompatibility of the device were also supported by consistent signaling during level walking at 14 days and 56 days post implantation (0.01 &plusmn; 0.007 mV, 0.012 &plusmn; 0.007 mV respectively; p &gt; 0.05) and the absence of histological inflammation. Collectively, we provide an updated workflow for the fabrication and testing of low-cost epimysial electrodes.

Author Spotlight: Epimysial Electrode Fabrication and Testing in ACL Injury Studies

Our purpose was to provide an updated, easy-to-follow guide on the fabrication and testing of epimysial electromyography electrodes. To that end, we provide instructions for material sourcing and a detailed walkthrough of the fabrication and testing process.

Electromyography (EMG) is a valuable diagnostic tool for detecting neuromuscular abnormalities. Implantable epimysial electrodes are commonly used to ...

In vivo Measurement of Knee Extensor Muscle Function in Mice

Skeletal muscle plasticity in response to countless conditions and stimuli mediates concurrent functional adaptation, both negative and positive. In the clinic and the research laboratory, maximal muscular strength is widely measured longitudinally in humans, with knee extensor musculature the most reported functional outcome. Pathology of the knee extensor muscle complex is well documented in aging, orthopedic injury, disease, and disuse; knee extensor strength is closely related to functional capacity and injury risk, underscoring the importance of reliable measurement of knee extensor strength. Repeatable, in vivo assessment of knee extensor strength in pre-clinical rodent studies offers valuable functional endpoints for studies exploring osteoarthritis or knee injury. We report an in vivo and non-invasive protocol to repeatedly measure isometric peak tetanic torque of the knee extensors in mice across time. We demonstrate consistency using this novel method to measure knee extensor strength with repeated assessment in multiple mice producing similar results.

Quantification of knee extensor maximal strength is imperative to understand functional adaptations to aging, disease, injury, and rehabilitation. We present a novel method to repeatedly measure in vivo knee extension isometric peak tetanic torque.

Skeletal muscle plasticity in response to countless conditions and stimuli mediates concurrent functional adaptation, both negative and positive. In the ...

In Vivo Measurement of Hindlimb Dorsiflexor Isometric Torque from Pig

Reliable assessment of skeletal muscle strength is arguably the most important outcome measure in neuromuscular and musculoskeletal disease and injury studies, particularly when evaluating regenerative therapies&#39; efficacy. Additionally, a critical aspect of translating many regenerative therapies is the demonstration of scalability and effectiveness in a large animal model. Various physiological preparations have been established to evaluate intrinsic muscle function properties in basic science studies, primarily in small animal models. The practices may be categorized as: in vitro (isolated fibers, fiber bundles, or whole muscle), in situ (muscle with intact vascularization and innervation but distal tendon attached to a force transducer), and in vivo (structures of the muscle or muscle unit remain intact). There are strengths and weaknesses to each of these preparations; however, a clear advantage of in vivo strength testing is the ability to perform repeated measurements in the same animal. Herein, the materials and methods to reliably assess isometric torque produced by the hindlimb dorsiflexor muscles in vivo in response to standard peroneal electrical stimulation in anesthetized pigs are presented.

In Vivo Measurement of Hindlimb Dorsiflexor Isometric Torque from Pig

The present protocol describes concise experimental details on the evaluation and interpretation of in vivo torque data obtained via electrical stimulation of the common peroneal nerve in anesthetized pigs.

Reliable assessment of skeletal muscle strength is arguably the most important outcome measure in neuromuscular and musculoskeletal disease and injury ...

Measuring Compound Muscle Action Potentials in Mouse Forelimb Muscles In Vivo

Source: Pollari, E. et al., <a href="https://app.jove.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 ...

Performing In Vivo and Ex Vivo Electrical Impedance Myography in Rodents

Summary

Explore More Videos

Chapters in this video

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

In Vitro Recording of Mesenteric Afferent Nerve Activity in Mouse Jejunal and Colonic Segments

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

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

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

Development of a Low-cost Epimysial Electromyography Electrode: A Simplified Workflow for Fabrication and Testing

In vivo Measurement of Knee Extensor Muscle Function in Mice

In Vivo Measurement of Hindlimb Dorsiflexor Isometric Torque from Pig

Measuring Compound Muscle Action Potentials in Mouse Forelimb Muscles In Vivo