Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

Summary

GABAergic presynaptic inhibition is a powerful inhibitory mechanism in the spinal cord important for motor and sensory signal integration in spinal cord networks. Underlying primary afferent depolarization can be measured by recording of dorsal root potentials (DRP). Here we demonstrate a method of in vivo recording of DRP in mice.

Abstract

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of primary afferent fibers mediated by GABAergic axo-axonal synapses (primary afferent depolarization). The strength of primary afferent depolarization can be measured by recording of volume-conducted potentials at the dorsal root (dorsal root potentials, DRP). Pathological changes of presynaptic inhibition are crucial in the abnormal central processing of certain pain conditions and in some disorders of motor hyperexcitability. Here, we describe a method of recording DRP in vivo in mice. The preparation of spinal cord dorsal roots in the anesthetized animal and the recording procedure using suction electrodes are explained. This method allows measuring GABAergic DRP and thereby estimating spinal presynaptic inhibition in the living mouse. In combination with transgenic mouse models, DRP recording may serve as a powerful tool to investigate disease-associated spinal pathophysiology. In vivo recording has several advantages compared to ex vivo isolated spinal cord preparations, e.g. the possibility of simultaneous recording or manipulation of supraspinal networks and induction of DRP by stimulation of peripheral nerves.

Introduction

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. It inhibits excitatory postsynaptic potentials (EPSPs) in monosynaptically excited motoneurons without changing the postsynaptic membrane potential and the excitability of the motoneurons^1-3. Primary afferent depolarization (PAD) induced by GABAergic axo-axonal synapses onto sensory presynaptic fibers is the underlying mechanism^4-7 (see also Figure1a). These synapses contain GABA_A- and GABA_B-receptors (GABA_AR and GABA_BR). GABA_AR activity leads to an increase in chloride conductance which elicits PAD due to the local ion distribution. This depolarization blocks the propagation of action potentials into the axon terminals and reduces their strength leading to a decreased Ca²⁺-influx and a reduction of transmitter release. Activation of GABA_B receptors does not contribute to PAD but leads to a reduction of Ca²⁺-influx thereby enhancing presynaptic inhibition. While the activation of GABA_AR seems to be involved in short term inhibition, GABA_BR are involved in long-term modulation^8-10. In addition to GABA, which accounts for the major part of PAD and presynaptic inhibition, other transmitters systems might also modulate and contribute to this mechanism^11,12.

Pathological changes in presynaptic inhibition seem to be crucial in several disease states e.g. peripheral inflammation and neuropathic pain^13,14, as well as abnormal central pain processing¹⁵, spinal cord injury¹⁶, and CNS disease with motor hyperexcitability mediated by defective GABAergic transmission^17,18. Thus, estimating presynaptic inhibition is worthwhile to investigate experimental pathological conditions on the spinal cord level in vivo. PAD gives rise to volume conducted potentials providing a direct measure of the presynaptic inhibition in the spinal cord. Those potentials are called dorsal root potentials (DRP) and can be measured from spinal cord dorsal roots after stimulation of adjacent dorsal roots⁷.

First measurements of DRP have been reported in cats and frogs¹⁹ and were intensively studied in cats by Eccles, Schmidt, and others in the early 1970s^3,4,20,21. While in vivo recordings of DRP in cats²² and rats²³ have been widely used, measurements in mice have been almost exclusively performed in ex vivo isolated spinal cord preparations^15,24. Here, we describe a method to record DRP in anesthetized mice in vivo allowing a direct measure of presynaptic inhibition in the intact organism.

Protocol

All experimental procedures mentioned in the following protocol were approved by the Thuringian state authorities (Thüringer Landesamt für Verbraucherschutz, Reg.-Nr. 02-044/12).

1. Preparations for Experiment

1. Fabrication of suction electrodes
   1. Pull a micropipette using a standard borosilicate glass capillary with a micropipette puller, e.g. a standard patch electrode.
   2. Brake the electrode to tip diameter of 0.5-1 mm (slightly larger than the diameter of the dorsal roots) using a diamond file.
   3. Heat polish the tip so it will not harm the dorsal root when it is sucked in. A standard lab torch will be suitable.
   4. Mount the glass filament on an electrode holder connected to a syringe through which negative pressure can be applied.
2. Preparation of solutions
   1. Injection anesthesia for mice: Mix 0.75 ml ketamine 10%, 0.24 ml xylazine 2% and 5 ml 0.9% saline. 10 µl/g bodyweight (BW) of the solution will be injected intraperitoneally (i.p.).
   2. Artificial cerebrospinal fluid (aCSF): Dilute in double distilled water the following salts (in mM): NaCl 134; KCl 3; KH₂PO₄ 1.25; MgSO₄•H₂O 2; NaHCO₃ 25; CaCl₂ 2; D-glucose 10. Add H₂O₂ to a final concentration of 0.003%. Use always freshly prepared solution.
3. Preparation of the recording setup (Figure 2)
   1. Connect the amplifier to a PC interface for digitizing data.
   2. Use a PC-based program or an analogue timer to trigger a square pulse stimulator and data acquisition on PC hard drive.
   3. Use chlorided silverwires connected to standard glass electrode holders as for stimulation and recording.
   4. Assemble three manipulators for stimulation, recording and reference electrode respectively around a stereotactic frame for mice, so that all electrodes have access to the prepared spinal cord later on.
   5. Connect tubings and syringes to the electrode holders to be apple to apply negative pressure.

2. General Comments for Animal Experiments and Animal Preparation for Recording Procedure

1. Perform all experiments using mice according to the guidelines of the respective institutional animal care and use committee. Perform all surgery and recordings under deep anesthesia ensuring that animal suffering is minimized.
2. Anesthetize the animal by i.p. injection of ketamine/xylazine (125mg and 8mg per g BW of ketamine and xylazine, respectively; 10 µl/g BW of the prepared solution as described above). If necessary during long-term recordings, additional injections can be made i.p. or i.m.
   Note: Repetitive i.m. injections of 0.05-0.1 ml of ketamine/xylazine solution in the upper thighs have proven to be suitable to keep the animal in deep anesthesia for up to 3 hr.
3. Use vet ointment on eyes to prevent dryness while under anesthesia.
4. Fix the head of the animal in a stereotactic frame and use a heating pad with rectal probe and reflex loop to control body temperature of the animal during the experiment. Fixation of the spinal cord in a stereotaxic frame is not necessary.
5. Before starting the preparation, check depth of narcosis by eyeblink reflex and twitching between toes of the mice. Reflexes should be abolished.
6. Open the skin along the midline above the spinal cord from the upper thoracic level to lower lumbar areas to get a clear operational field using a scalpel. Do not use scissors to make skin incisions. Carefully loose the skin from the underlying tissue. During subsequent steps keep the wound moistened by 0.9% saline.
7. Cut tendons and connective tissue on both side of the vertebrae from lumbar to thoracic levels using scalpel and scissor.
8. Remove the spinous processes and rest of connective tissue around the vertebrae using a small nipper.
9. Carefully crack vertebrae with the nipper starting from lumbar levels (L4/L5) under a dissecting microscope. Shove the tip of the nipper in the space between the vertebrae and the spinal cord and lift bone pieces apart. Do not harm the dura and avoid pressure to the spinal cord. Both are critical for success. Keep the spinal cord moistened during the whole procedure. Proceed to mid-thoracic levels.

3. Separating of Dorsal Roots and DRP Recording (Figure 2)

1. Open the dura mater carefully using a thin needle (30 G) with a bent tip. Use aCSF for moistening from now on.
2. Separate the dorsal roots as far as possible using the gauge and cut two adjacent roots as distal as possible. Vigorous pulling on the roots during separation affects the success of the experiment.
3. Lower the suction electrode down to the dorsal roots. Add as much aCSF to the spinal cord as possible because liquid helps to suck in the dorsal roots.
4. Suck the cut end of one dorsal root into one glass pipettes by applying negative pressure through a syringe. If needed, move the dorsal root in front of the electrode opening using a fine needle.
5. If the dorsal root lies dry within the suction electrode, add some aCSF to its tip while carefully sucking until the pipette is sufficiently filled with aCSF.
6. After the dorsal root is sucked in, raise the electrode tip from the spinal cord. Take care that no “water bridge” between the pipette tip and the spinal cord short-circuit the recording/stimulation electrode.
7. Repeat steps 3.3-3.6 for an ipsilateral adjacent root.
8. Position the reference electrode as close as possible to the dorsal roots and keep it moistened by applying aCSF.
9. By establishing the recording setup, one root is already chosen for stimulation, the other for recording. Increase voltage stepwise while recording from the second dorsal root is done in current clamp mode. One may notice a short downward deflection which is followed by a slow, long-lasting upward deflection representing the DRP.
10. Adjust the stimulus voltage to supramaximal levels and record several sweeps (at least 20 - 30 sweeps/dorsal root at 0.1 Hz, filter between 0.3 Hz-3 kHz).
11. Record short trains of three stimuli (100 Hz) to get information about the time-dependent summation of the DRP.
12. Switch recording and stimulation site for additional contralateral recordings.
13. Animals are not intended to survive the procedure. Sacrifice animals after last recording by decapitation while still under deep anesthesia (ketamine/xylazine, see above, step 2.2).

4. Data Analysis

1. Transfer data to analysis program (e.g. Sigma Plot, Igor Pro, or MATLAB).
2. Calculate average traces from 20-30 sweeps.
3. Take the maximal amplitude of the voltage deflection (from baseline; DRP peak amplitude) for further analysis (Figure 3).
4. Calculate the ratio of the DRP peak amplitude after three pulses and single pulse to gain a measure of the time dependent summation of subsequent potentials.

Results

Typical DRP traces are shown in Figure 3. The prominent stimulation artifact is usually followed by a short downward deflection. Thereafter a slow, long-lasting upward deflection, representing the DRP is clearly distinguishable. In a subset of recordings, dorsal root reflexes are visible as small spikes on top of the DRP. In normal wild-type mice, dorsal root reflexes appear most often when stimulation voltage is excessive. As the dorsal root reflexes cannot be elicited with a high reproducibility in thi...

Discussion

Extra- and intracellular electrophysiological recordings of neuronal activity and synaptic potentials in vivo are state of the art techniques in investigating CNS neuronal functions and pathophysiology. Spinal integration is critical for motor function, e.g. limb movement and for multimodal sensory perception. Presynaptic inhibition is one critical mechanism in this computational process ensuring appropriate responses to sensory inputs. GABAergic synapses on Ia afferent fibers inhibit the excitation of ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Glass tubing (inner diameter 1.16 mm)	Science Products (Hofheim, Germany)	GB200F-10	Other glass tubing might also be suitable
Superfusion solution (sterile, 0,9% NaCl)	Braun Melsungen AG	3570350
Rompun 2% (Xylazine)	Bayer Animal Health GmbH (Leverkusen, Germany)
Ketamine 10%	Medistar GmbH (Ascheberg, Germany)	KETAMIN 10%
30 G Microneedle/ Sterican	Braun Melsungen AG	4656300
Salts for aCSF	Sigma-Aldrich	Diverse
S88 Dual Output Square Pulse	Grass Technologies (Warwick, USA)	S88X
SIU5 RF Transformer Isolation Unit	Grass Technologies (Warwick, USA)	SIU-V
InstruTECH LIH 8+8	HEKA (Lambrecht, Deutschland)	LIH 8+8 + Patchmaster software
Universal amplifier	NPI (Tamm, Deutschland)	ELC-03X
Micropipette puller	Sutter Instruments (Novato, USA)	P-1000
Dissecting microscope	Olympus (Tokyo, Japan)
Micromanipulator	Sutter Instruments (Novato, USA)	MPC-200/MPC-325	Mechanical micromanipulators also possible
Homeothermic Blanket System	Stoelting (Wood Dale, USA)	50300V
Intra-/extracellular recording electrode holder	Harvard Apparatus (Holliston, USA)	641227