Performing In Vivo and Ex Vivo Electrical Impedance Myography in Rodents

Summary

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).

Abstract

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.

Introduction

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 therapy^1,2,3. 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 "chargeability" of the tissue, a measure related to the tissue's ability to store charge (capacitance). From the reactance and resistance, the phase angle (θ) is calculated via the following equation: , 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 them^4,5. 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 "neuromuscular disorders" 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 aging^10,11. 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 size^12,13, inflammatory changes and edema¹⁴, and connective tissue and fat content^15,16.

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 ultrasound^18,19. 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.

Protocol

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

1. Place the animal in an anesthesia box to induce anesthesia.
   NOTE: For rats, 1.5%-3.5% isoflurane and 2 O₂ L·min^-1 were used, and for mice, 2% isoflurane and 1 O₂ L·min^-1 were used.
2. 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·min^-1.
3. Place the animal's leg to be analyzed at a 45° angle with the hip joint (knee extended) and secure the foot with medical tape.
4. Use a hair clipper to trim the fur overlaying the gastrocnemius muscle.
5. Apply a thick layer of depilatory cream over the animal'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.
6. Connect the surface array (Figure 1) to the EIM device and let the electrodes rest on a piece of gauze soaked in saline solution.
7. Place the surface array directly on the skin over the gastrocnemius muscle, oriented longitudinally to the muscle fibers.
8. 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.
9. Rotate the surface array by 90° and reposition it on the skin over the gastrocnemius to obtain the transverse measurements (check for green bars indicating the stability).
10. 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.

2. In vivo needle array EIM

1. 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.
2. 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).
3. 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.
4. Gently remove the array and reinsert it through the skin and into the muscle at a 90° 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.

3. Ex vivo EIM

1. 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 ± 25 Ω over the frequency range from 1 kHz to 1 MHz.
2. Euthanize the animal according to respective IACUC guidelines.
3. 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.
4. 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.
5. 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.
6. 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.
7. 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.
8. 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.
9. Open the dielectric cell and reorient the muscle in the transverse direction by rotating it 90°. Reattach the top of the dielectric cell. Acquire the transverse measurement.

Results

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 ...

Discussion

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 ...

Materials

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