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

Summary

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.

Abstract

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 vivo^2-4 , including our team who published a new preparation which allowed us to obtain very stable recordings of motoneurons in vivo in adult mice^5,6. 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 circuitry^7-9, or reproducting human disease^10,11, we expect this technique to be an essential tool for the study of spinal motor system.

Protocol

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.

1. Cover the snout of the mouse with a mask delivering pure O₂ at a flow around 100 ml/min.
2. Using a hair clipper, shave the dorsal region from the nape of the neck to the base of the tail.
3. Shave also the right hindlimb.
4. Flip mouse to supine position but take care to leave the oxygen mask in place.

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 °C and 38 °C.

6. Tracheotomy and Artificial Ventilation

1. Using blunt scissors, make a cut over the trachea and pull the skin away on both sides.
2. Using blunt forceps, tear apart the salivary gland so as to expose two thin muscles (Sternohyoid) covering the trachea.
3. Using blunt forceps, separate the two muscles along their medial separation to reveal the trachea.
4. Using a Dumont 7 forceps, slide two length of 4.0 silk suture under the trachea.
5. Make a transversal cut in the trachea in between two cartilaginous rings but take care not to completely section the trachea.
6. 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 (μcapstar, 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 μl) so that the mouse is not fighting against the artificial ventilation and the end-tidal pCO₂ is stable between 4 and 5%.

7. Placement of the Intravenous Lines

1. 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.
2. Do the following steps twice, one set for each of these trunks:
   1. Using Dumont 4 or 5 forceps, carefully separate the vein from its surrounding conjunctive tissue.
   2. 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.
   3. 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.
   4. Using very fine iris scissors, make a small transverse incision in the vein, taking great care not to section the vein completely.
   5. Insert a pre-filled 1Fr catheter (premicath, Vygon) in the opening, up to the vessel clip.
   6. 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.
   7. Secure the catheter by tying it with both sutures on both sides of the insertion.
3. 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 μg/kg/min of ketamine/xylazine¹². The other catheter is connected to a syringe pump for a slow intravenous infusion (50 μl/hr) of a 4 % glucose solution containing NaHCO₃ (1 %) and plasmion (14 %).

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

1. 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.
2. 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.
3. 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.
4. Dissect out the Sural nerve.
5. 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.
6. 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).
7. 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.
8. Cover the whole hindlimb with a gauze imbibed with saline to prevent dessication while proceeding with the next step.

10. Laminectomy

1. Using scissors, make an incision along the backbone. Separate the skin from the underlying muscles.
2. 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.
3. 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.
4. 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.
5. Using fine rongeurs, remove the spinal processes then the laminae over T13 and L1, thereby exposing the spinal cord.
6. Cover the exposed spinal cord with small bits of cotton or spongel imbibed with saline to prevent desiccation.
7. 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).
8. Once the Kwik-Cast has dried, remove the cotton covering the spinal cord, and fill the bath with mineral oil.
9. 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.
10. Lower the raised platform on which the mouse is lying so as to keep it suspended by the vertebral clamp.
11. Use another vertebral clamp to clamp a spinous process of the sacral region to support the back region of the animal.
12. A third clamp is used to immobilize the right hindlimb and ankle, flexed at a 90° angle at the knee.

11. Achilles Tendon Dissection

1. With the limb bent at 90° 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.
2. Cut the tendon of the Plantaris muscle close to the calcaneus, then cut it again to remove the tendon completely.
3. Using a threaded needle, attach 6/0 silk thread through the Achilles tendon and make a triple knot around the tendon.
4. 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.
5. 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.
6. Place the Common Peroneal nerve and the Tibial nerve on two bipolar hook electrodes.
7. Place the Triceps Surae nerve on the cathode of a hook electrode, with the anode touching a nearby muscle.
8. Connect all stimulating electrodes to an isolation unit.
9. 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.

12. Step Twelve

Stimulate the Triceps Surae nerve using a 50 μsec square pulse of increasing intensity at a low frequency (<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.

1. Pull a glass micropipette to a ~1 μm 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 MΩ).
2. 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.
3. 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.

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.

Results

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...

Discussion

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...

Materials

List of materials used in this article

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