We thank Manfred Heckmann for helpful discussions during establishing of the method. Further, we thank Claudia Sommer for technical assistance and Frank Schubert for support producing the video. The work was supported by the Federal Ministry of Education and Research (BMBF), Germany, FKZ: 01EO1002 and the Interdisciplinary Center for Clinical Research (IZKF) of Jena University Hospital.

All experimental procedures mentioned in the following protocol were approved by the Thuringian state authorities (Th&#252;ringer Landesamt f&#252;r Verbraucherschutz, Reg.-Nr. 02-044/12).
1. Preparations for Experiment
<ol>
	<li>Fabrication of suction electrodes
	<ol>
		<li>Pull a micropipette using a standard borosilicate glass capillary with a micropipette puller, e.g. a standard patch electrode.</li>
		<li>Brake the electrode to tip diameter of 0.5-1 mm (slightly larger than the diameter of the dorsal roots) using a diamond file.</li>
		<li>Heat polish the tip so it will not harm the dorsal root when it is sucked in. A standard lab torch will be suitable.</li>
		<li>Mount the glass filament on an electrode holder connected to a syringe through which negative pressure can be applied.</li>
	</ol>
	</li>
	<li>Preparation of solutions
	<ol>
		<li>Injection anesthesia for mice: Mix 0.75 ml ketamine 10%, 0.24 ml xylazine 2% and 5 ml 0.9% saline. 10 &#181;l/g bodyweight (BW) of the solution will be injected intraperitoneally (i.p.).</li>
		<li>Artificial cerebrospinal fluid (aCSF): Dilute in double distilled water the following salts (in mM): NaCl 134; KCl 3; KH2PO4 1.25; MgSO4&#8226;H2O 2; NaHCO3 25; CaCl2 2; D-glucose 10. Add H2O2 to a final concentration of 0.003%. Use always freshly prepared solution.</li>
	</ol>
	</li>
	<li>Preparation of the recording setup (Figure 2)
	<ol>
		<li>Connect the amplifier to a PC interface for digitizing data.</li>
		<li>Use a PC-based program or an analogue timer to trigger a square pulse stimulator and data acquisition on PC hard drive.</li>
		<li>Use chlorided silverwires connected to standard glass electrode holders as for stimulation and recording.</li>
		<li>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.</li>
		<li>Connect tubings and syringes to the electrode holders to be apple to apply negative pressure.</li>
	</ol>
	</li>
</ol>
2. General Comments for Animal Experiments and Animal Preparation for Recording Procedure
<ol>
	<li>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.</li>
	<li>Anesthetize the animal by i.p. injection of ketamine/xylazine (125mg and 8mg per g BW of ketamine and xylazine, respectively; 10 &#181;l/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.</li>
	<li>Use vet ointment on eyes to prevent dryness while under anesthesia.</li>
	<li>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.</li>
	<li>Before starting the preparation, check depth of narcosis by eyeblink reflex and twitching between toes of the mice. Reflexes should be abolished.</li>
	<li>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.</li>
	<li>Cut tendons and connective tissue on both side of the vertebrae from lumbar to thoracic levels using scalpel and scissor.</li>
	<li>Remove the spinous processes and rest of connective tissue around the vertebrae using a small nipper.</li>
	<li>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.</li>
</ol>
3. Separating of Dorsal Roots and DRP Recording (Figure 2)
<ol>
	<li>Open the dura mater carefully using a thin needle (30 G) with a bent tip. Use aCSF for moistening from now on.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>After the dorsal root is sucked in, raise the electrode tip from the spinal cord. Take care that no &#8220;water bridge&#8221; between the pipette tip and the spinal cord short-circuit the recording/stimulation electrode.</li>
	<li>Repeat steps 3.3-3.6 for an ipsilateral adjacent root.</li>
	<li>Position the reference electrode as close as possible to the dorsal roots and keep it moistened by applying aCSF.</li>
	<li>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.</li>
	<li>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).</li>
	<li>Record short trains of three stimuli (100 Hz) to get information about the time-dependent summation of the DRP.</li>
	<li>Switch recording and stimulation site for additional contralateral recordings.</li>
	<li>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).</li>
</ol>
4. Data Analysis
<ol>
	<li>Transfer data to analysis program (e.g. Sigma Plot, Igor Pro, or MATLAB).</li>
	<li>Calculate average traces from 20-30 sweeps.</li>
	<li>Take the maximal amplitude of the voltage deflection (from baseline; DRP peak amplitude) for further analysis (Figure 3).</li>
	<li>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.</li>
</ol>

<ol>
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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Depolarization+of+central+terminals+of+Group+I+afferent+fibres+from+muscle.">Depolarization of central terminals of Group I afferent fibres from muscle.</a> J. Physiol. 160, 62-93 (1962).</li><li>Eccles, J. C., Schmidt, R., Willis, W. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pharmacological+Studies+on+Presynaptic+Inhibition.">Pharmacological Studies on Presynaptic Inhibition.</a> J. Physiol. 168, 500-530 (1963).</li><li>Maxwell, D. J., Bannatyne, B. 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D., Lidierth, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Five+sources+of+a+dorsal+root+potential:+their+interactions+and+origins+in+the+superficial+dorsal+horn.">Five sources of a dorsal root potential: their interactions and origins in the superficial dorsal horn.</a> J. Neurophysiol. 78, 860-871 (1997).</li><li>Rudomin, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+search+of+lost+presynaptic+inhibition.">In search of lost presynaptic inhibition.</a> Exp. Brain Res. 196, 139-151 (2009).</li><li>Rudomin, P., Schmidt, R. 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Lett. 217, 153-156 (1996).</li><li>Enriquez-Denton, M., Manjarrez, E., Rudomin, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistence+of+PAD+and+presynaptic+inhibition+of+muscle+spindle+afferents+after+peripheral+nerve+crush.">Persistence of PAD and presynaptic inhibition of muscle spindle afferents after peripheral nerve crush.</a> Brain Res. 1027, 179-187 (2004).</li><li>Wall, P. D., Devor, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+peripheral+nerve+injury+on+dorsal+root+potentials+and+on+transmission+of+afferent+signals+into+the+spinal+cord.">The effect of peripheral nerve injury on dorsal root potentials and on transmission of afferent signals into the spinal cord.</a> Brain Res. 209, 95-111 (1981).</li><li>Witschi, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Presynaptic+&#945;2-GABAA+Receptors+in+Primary+Afferent+Depolarization+and+Spinal+Pain+Control.">Presynaptic &#945;2-GABAA Receptors in Primary Afferent Depolarization and Spinal Pain Control.</a> J. Neurosci. 31, 8134-8142 (2011).</li><li>Calancie, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+that+alterations+in+presynaptic+inhibition+contribute+to+segmental+hypo-+and+hyperexcitability+after+spinal+cord+injury+in.">Evidence that alterations in presynaptic inhibition contribute to segmental hypo- and hyperexcitability after spinal cord injury in.</a> 89, 177-186 (1993).</li><li>Geis, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stiff+person+syndrome-associated+autoantibodies+to+amphiphysin+mediate+reduced+GABAergic+inhibition.">Stiff person syndrome-associated autoantibodies to amphiphysin mediate reduced GABAergic inhibition.</a> Brain. 133, 3166-3180 (2010).</li><li>Geis, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+IgG+directed+against+amphiphysin+induces+anxiety+behavior+in+a+rat+model+after+intrathecal+passive+transfer.">Human IgG directed against amphiphysin induces anxiety behavior in a rat model after intrathecal passive transfer.</a> J. Neural Transm. 119 (8), 981-985 (2012).</li><li>Barron, D. H., Matthews, B. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+interpretation+of+potential+changes+in+the+spinal+cord.">The interpretation of potential changes in the spinal cord.</a> J. Physiol. 92, 276-321 (1938).</li><li>Schmidt, R. F., Trautwein, W., Zimmermann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dorsal+root+potentials+evoked+by+natural+stimulation+of+cutaneous+afferents.">Dorsal root potentials evoked by natural stimulation of cutaneous afferents.</a> Nature. 212, 522-523 (1966).</li><li>Eccles, J. C., Schmidt, R. F., Willis, W. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Presynaptic+inhibition+of+the+spinal+monosynaptic+reflex+pathway.">Presynaptic inhibition of the spinal monosynaptic reflex pathway.</a> J. Physiol. 161, 282-297 (1962).</li><li>Manjarrez, E., Rojas-Piloni, J. G., Jimenez, I., Rudomin, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulation+of+synaptic+transmission+from+segmental+afferents+by+spontaneous+activity+of+dorsal+horn+spinal+neurones+in+the+cat.">Modulation of synaptic transmission from segmental afferents by spontaneous activity of dorsal horn spinal neurones in the cat.</a> J. 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The authors declare no competing financial interests.

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 ...

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 motoneurons1-3. Primary afferent depolarization (PAD) induced by GABAergic axo-axonal synapses onto sensory presynaptic fibers is the underlying mechanism4-7 (see also Figure1a). These synapses contain GABAA- and GABAB-receptors (GABAAR and GABABR). GABAAR 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 GABAB receptors does not contribute to PAD but leads to a reduction of Ca2+-influx thereby enhancing presynaptic inhibition. While the activation of GABAAR seems to be involved in short term inhibition, GABABR are involved in long-term modulation8-10. 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 mechanism11,12.
Pathological changes in presynaptic inhibition seem to be crucial in several disease states e.g. peripheral inflammation and neuropathic pain13,14, as well as abnormal central pain processing15, spinal cord injury16, and CNS disease with motor hyperexcitability mediated by defective GABAergic transmission17,18. 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 1970s3,4,20,21. 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 preparations15,24. Here, we describe a method to record DRP in anesthetized mice in vivo allowing a direct measure of presynaptic inhibition in the intact organism.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Glass tubing (inner diameter 1.16 mm)</td>
			<td>Science Products (Hofheim, Germany)</td>
			<td>GB200F-10</td>
			<td>Other glass tubing might also be suitable</td>
		</tr>
		<tr>
			<td>Superfusion solution (sterile, 0,9% NaCl)</td>
			<td>Braun Melsungen AG&#160;</td>
			<td>3570350</td>
			<td></td>
		</tr>
		<tr>
			<td>(Melsungen, Germany)</td>
		</tr>
		<tr>
			<td>Rompun 2% (Xylazine)</td>
			<td>Bayer Animal Health GmbH (Leverkusen, Germany)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamin 10%</td>
			<td>Medistar GmbH (Ascheberg, Germany)</td>
			<td>KETAMIN 10%</td>
			<td></td>
		</tr>
		<tr>
			<td>30G micro needle/ Sterican</td>
			<td>Braun Melsungen AG&#160;</td>
			<td>4656300</td>
			<td></td>
		</tr>
		<tr>
			<td>(Melsungen, Geramny)</td>
		</tr>
		<tr>
			<td>Salts for aCSF</td>
			<td>Sigma-Aldrich&#160;</td>
			<td>Diverse</td>
			<td></td>
		</tr>
		<tr>
			<td>S88 Dual Output Square Pulse</td>
			<td>Grass Technologies (Warwick, USA)</td>
			<td>S88X</td>
			<td></td>
		</tr>
		<tr>
			<td>Stimulator</td>
		</tr>
		<tr>
			<td>SIU5 RF Transformer Isolation Unit</td>
			<td>Grass Technologies (Warwick, USA)</td>
			<td>SIU-V</td>
			<td></td>
		</tr>
		<tr>
			<td>InstruTECH LIH 8+8</td>
			<td>HEKA (Lambrecht, Deutschland)</td>
			<td>LIH 8+8 + Patchmaster software</td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquisition&#160;</td>
		</tr>
		<tr>
			<td>Universal amplifier</td>
			<td>npi (Tamm, Deutschland)</td>
			<td>ELC-03X</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instruments (Novato, USA)</td>
			<td>P-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>Olympus (Tokyo, Japan)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Sutter Instruments (Novato, USA)</td>
			<td>MPC-200/MPC-325</td>
			<td>Mechanical micromanipulators also possible</td>
		</tr>
		<tr>
			<td>Homeothermic Blanket System</td>
			<td>Stoelting (Wood Dale, USA)</td>
			<td>50300V</td>
			<td></td>
		</tr>
		<tr>
			<td>Intra-/extracellular recording electrode holder</td>
			<td>Harvard Apparatus (Holliston, USA)</td>
			<td>641227</td>
			<td></td>
		</tr>
	</tbody>
</table>

measuring spinal presynaptic inhibition in mice by dorsal root potential recording in vivo

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.

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...

Watch this Scientific Journal Video about Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo at JoVE.com

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

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.

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

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of ...

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13.9K Views.  Jena University Hospital, Jena, Germany. 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.

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

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These neurotropic cancer cells have a unique ability to migrate unidirectionally along nerves towards the central nervous system (CNS). The dorsal root ganglia (DRG)/cancer cell model is a three dimensional (3D) in vitro model frequently used for studying the interaction between neural stroma and cancer cells. In this model, mouse or human cancer cell lines are grown in ECM adjacent to preparations of freshly dissociated cultured DRG. In this article, the DRG isolation protocol from mice, and implantation in petri dishes for co-culturing with pancreatic cancer cells are demonstrated. Five days after implantation, the cancer cells made contact with the DRG neurites. Later, these cells formed bridgeheads to facilitate more extensive polarized, neurotropic migration of cancer cells.

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

This video article shows the use of the dorsal root ganglia (DRG)/cancer cell model in pancreatic ductal adenocarcinoma.

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These ...

Polygraphic Recording Procedure for Measuring Sleep in Mice

Recording of the epidural electroencephalogram (EEG) and electromyogram (EMG) in small animals, like mice and rats, has been pivotal to study the homeodynamics and circuitry of sleep-wake regulation. In many laboratories, a cable-based sleep recording system is used to monitor the EEG and EMG in freely behaving mice in combination with computer software for automatic scoring of the vigilance states on the basis of power spectrum analysis of EEG data. A description of this system is detailed herein. Steel screws are implanted over the frontal cortical area and the parietal area of 1 hemisphere for monitoring EEG signals. In addition, EMG activity is monitored by the bilateral placement of wires in both neck muscles. Non-rapid eye movement (Non-REM; NREM) sleep is characterized by large, slow brain waves with delta activity below 4 Hz in the EEG, whereas a shift from low-frequency delta activity to a rapid low-voltage EEG in the theta range between 6 and 10 Hz can be observed at the transition from NREM to REM sleep. By contrast, wakefulness is identified by low- to moderate-voltage brain waves in the EEG trace and significant EMG activity.

The recording of electroencephalogram (EEG) and electromyogram (EMG) in freely behaving mice is a critical step to correlate behavior and physiology with sleep and wakefulness. The experimental protocol described herein provides a cable-based system for acquiring EEG and EMG recordings in mice.

Recording of the epidural electroencephalogram (EEG) and electromyogram (EMG) in small animals, like mice and rats, has been pivotal to study the ...

Hydraulic Extrusion of the Spinal Cord and Isolation of Dorsal Root Ganglia in Rodents

Traditionally, the spinal cord is isolated by laminectomy, i.e. by breaking open the spinal vertebrae one at a time. This is both time consuming and may result in damage to the spinal cord caused by the dissection process. Here, we show how the spinal cord can be extruded using hydraulic pressure. Handling time is significantly reduced to only a few minutes, likely decreasing protein damage. The low risk of damage to the spinal cord tissue improves subsequent immunohistochemical analysis. By performing hydraulic spinal cord extrusion instead of traditional laminectomy, the rodents can further be used for DRG isolation, thereby lowering the number of animals and allowing analysis across tissues from the same rodent. We demonstrate a consistent method to identify and isolate the DRGs according to their localization relative to the costae. It is, however, important to adjust this method to the particular animal used, as the number of spinal cord segments, both thoracic and lumbar, may vary according to animal type and strain. In addition, we illustrate further processing examples of the isolated tissues.

Here, we present a protocol for hydraulic extrusion of the spinal cord as well as identification and isolation of specific dorsal root ganglia (DRGs) in the same rodent. Compared to standard spinal cord isolation methods, this method is significantly faster and reduces the risk of tissue damage.

Traditionally, the spinal cord is isolated by laminectomy, i.e. by breaking open the spinal vertebrae one at a time. This is both time consuming and may ...

Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each other anatomically, it is important to identify an appropriate model to use for the study of axon regeneration. By using a suitable model, we can formulate a specific treatment based on the severity of injury, the neuronal cell type of interest, and the desired spinal tract for assessing regeneration. Within the sensory pathway, DRG neurons are responsible for relaying sensory information from the periphery to the CNS. We present here a protocol that uses a DRG injection with a viral vector and a concurrent dorsal root crush injury in the lower cervical spinal cord of an adult rat as a model to study sensory axon regeneration. As demonstrated using a control virus, AAV5-GFP, we show the effectiveness of a direct DRG injection in transducing DRG neurons and tracing sensory axons into the spinal cord. We also show the effectiveness of the dorsal root crush injury in denervating the forepaw as an injury model for evaluating axon regeneration. Despite the requirement for specialized training to perform this invasive surgical procedure, the protocol is flexible, and potential users can modify many parts to accommodate their experimental requirements. Importantly, it can serve as a foundation for those in search of a suitable animal model for their studies. We believe that this article will help new users to learn the procedure in a very efficient and effective manner.

This protocol presents the use of a dorsal root ganglion (DRG) injection with a viral vector and a concurrent dorsal root crush injury in an adult rat as a model to study sensory axon regeneration. This model is suitable for investigating the use of gene therapy to promote sensory axon regeneration.

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each ...

Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive targets for injection of novel therapeutics aimed at treating chronic pain. In small animal models, laminectomy has been used to facilitate DRG injection because it involves surgical removal of the vertebral bone surrounding each DRG. We demonstrate a technique for intraganglionic injection of lumbar DRG in a large animal species, namely, swine. Laminotomy is performed to allow direct access to DRG using standard neurosurgical techniques, instruments, and materials. Compared with more extensive bone removal via laminectomy, we implement laminotomy to conserve spinal anatomy while achieving sufficient DRG access. Intraoperative progress of DRG injection is monitored using a non-toxic dye. Following euthanasia on post-operative day 21, the success of injection is determined by histology for intraganglionic distribution of 4&#39;,6-diamidino-2-phenylindole (DAPI). We inject a biologically inactive solution to demonstrate the protocol. This method could be applied in future preclinical studies to target therapeutic solutions to DRG. Our methodology should facilitate testing the translatability of intraganglionic small animal paradigms in a large animal species. Additionally, this protocol may serve as a key resource for those planning preclinical studies of DRG injection in swine.

We describe a method for laminotomy in swine that provides access to lumbar dorsal root ganglia (DRG) for intraganglionic injection. Injection progress is monitored intraoperatively and histologically confirmed up to 21 days after surgery. This protocol could be used for future preclinical studies involving DRG injection.

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for nerve fibers in the central and peripheral nervous systems. This method is particularly applicable to dorsal root ganglia (DRG), which are primary sensory neurons that exhibit a pseudo-unipolar structure of nervous processes. The patterns and features of the action potentials passed along axons are recordable in these neurons. The present study uses in vivo single-fiber recordings to observe the conduction failure of sciatic nerves in complete Freund&#8217;s adjuvant (CFA)-treated rats. As the underlying mechanism cannot be studied using in vivo single-fiber recordings, patch-clamp-recordings of DRG neurons are performed on preparations of intact DRG with the attached sciatic nerve. These recordings reveal a positive correlation between conduction failure and the rising slope of the after-hyperpolarization potential (AHP) of DRG neurons in CFA-treated animals. The protocol for in vivo single fiber-recordings allows the classification of nerve fibers via the measurement of conduction velocity and monitoring of abnormal conditions in nerve fibers in certain diseases. Intact DRG with attached peripheral nerve allows observation of the activity of DRG neurons in most physiological conditions. Conclusively, single-fiber recording combined with electrophysiological recording of intact DRGs is an effective method to examine the role of conduction failure during the analgesic process.

Single-fiber recording is an effective electrophysiological technique that is applicable to the central and peripheral nervous systems. Along with the preparation of intact DRG with the attached sciatic nerve, the mechanism of conduction failure is examined. Both protocols improve the understanding of the peripheral nervous system&#39;s relationship with pain.

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for ...