This work was made possible thanks to financial support from the Fondation pour la Recherche M&eacute;dicale (FRM), the Milton Safenowitz PostDoctoral Fellowship for ALS Research (ALS Association), NIH NINDS Grants NS05462 and NS034382, and ANR Grant HyperMND.

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.
<ol>
	<li>Cover the snout of the mouse with a mask delivering pure O2 at a flow around 100 ml/min.</li>
	<li>Using a hair clipper, shave the dorsal region from the nape of the neck to the base of the tail.</li>
	<li>Shave also the right hindlimb.</li>
	<li>Flip mouse to supine position but take care to leave the oxygen mask in place.</li>
</ol>
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 &deg;C and 38 &deg;C.
6. Tracheotomy and Artificial Ventilation
<ol>
	<li>Using blunt scissors, make a cut over the trachea and pull the skin away on both sides.</li>
	<li>Using blunt forceps, tear apart the salivary gland so as to expose two thin muscles (Sternohyoid) covering the trachea.</li>
	<li>Using blunt forceps, separate the two muscles along their medial separation to reveal the trachea.</li>
	<li>Using a Dumont 7 forceps, slide two length of 4.0 silk suture under the trachea.</li>
	<li>Make a transversal cut in the trachea in between two cartilaginous rings but take care not to completely section the trachea.</li>
	<li>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 (&mu;capstar, 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 &mu;l) so that the mouse is not fighting against the artificial ventilation and the end-tidal pCO2 is stable between 4 and 5%.</li>
</ol>
7. Placement of the Intravenous Lines
<ol>
	<li>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.</li>
	<li>Do the following steps twice, one set for each of these trunks:
	<ol>
		<li>Using Dumont 4 or 5 forceps, carefully separate the vein from its surrounding conjunctive tissue.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Using very fine iris scissors, make a small transverse incision in the vein, taking great care not to section the vein completely.</li>
		<li>Insert a pre-filled 1Fr catheter (premicath, Vygon) in the opening, up to the vessel clip.</li>
		<li>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.</li>
		<li>Secure the catheter by tying it with both sutures on both sides of the insertion.</li>
	</ol>
	</li>
	<li>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 &mu;g/kg/min of ketamine/xylazine12. The other catheter is connected to a syringe pump for a slow intravenous infusion (50 &mu;l/hr) of a 4 % glucose solution containing NaHCO3 (1 %) and plasmion (14 %).</li>
</ol>
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
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Dissect out the Sural nerve.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>Cover the whole hindlimb with a gauze imbibed with saline to prevent dessication while proceeding with the next step.</li>
</ol>
10. Laminectomy
<ol>
	<li>Using scissors, make an incision along the backbone. Separate the skin from the underlying muscles.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Using fine rongeurs, remove the spinal processes then the laminae over T13 and L1, thereby exposing the spinal cord.</li>
	<li>Cover the exposed spinal cord with small bits of cotton or spongel imbibed with saline to prevent desiccation.</li>
	<li>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).</li>
	<li>Once the Kwik-Cast has dried, remove the cotton covering the spinal cord, and fill the bath with mineral oil.</li>
	<li>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.</li>
	<li>Lower the raised platform on which the mouse is lying so as to keep it suspended by the vertebral clamp.</li>
	<li>Use another vertebral clamp to clamp a spinous process of the sacral region to support the back region of the animal.</li>
	<li>A third clamp is used to immobilize the right hindlimb and ankle, flexed at a 90&deg; angle at the knee.</li>
</ol>
11. Achilles Tendon Dissection
<ol>
	<li>With the limb bent at 90&deg; 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.</li>
	<li>Cut the tendon of the Plantaris muscle close to the calcaneus, then cut it again to remove the tendon completely.</li>
	<li>Using a threaded needle, attach 6/0 silk thread through the Achilles tendon and make a triple knot around the tendon.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Place the Common Peroneal nerve and the Tibial nerve on two bipolar hook electrodes.</li>
	<li>Place the Triceps Surae nerve on the cathode of a hook electrode, with the anode touching a nearby muscle.</li>
	<li>Connect all stimulating electrodes to an isolation unit.</li>
	<li>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.</li>
</ol>
12. Step Twelve
Stimulate the Triceps Surae nerve using a 50 &mu;sec square pulse of increasing intensity at a low frequency (&lt;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.
<ol>
	<li>Pull a glass micropipette to a ~1 &mu;m 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 M&Omega;).</li>
	<li>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.</li>
	<li>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.</li>
</ol>
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.

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No conflicts of interest declared.

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...

A correction was made to <a href="http://www.jove.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>. There was an error in the name of one author, Marin Manuel. The author's name has been corrected to:
Marin Manuel 
 
instead of: 
 
Manuel Marin

A correction to author list has been made for article <a href="http://www.jove.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>.

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

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Atropine sulfate</td>
 <td>Aguettant</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Methylprenidsolone</td>
 <td>Pfizer</td>
 <td>Solu-Medrol</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium pentobarbitone</td>
 <td>Sanofi-Aventis</td>
 <td>Pentobarbital</td>
 <td></td>
 </tr>
 <tr>
 <td>Ketamine</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Xylazine</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Glucose</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Plasma expander</td>
 <td>Roger Bellon</td>
 <td>Plasmagel</td>
 <td></td>
 </tr>
 <tr>
 <td>Blunt scissors</td>
 <td>FST</td>
 <td>14079-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Blunt fine scissors</td>
 <td>FST</td>
 <td>15025-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Vannas Spring Scissors</td>
 <td>FST</td>
 <td>15002-08</td>
 <td></td>
 </tr>
 <tr>
 <td>Fine forceps serrated</td>
 <td>FST</td>
 <td>11370-32</td>
 <td></td>
 </tr>
 <tr>
 <td>Fine forceps serrated</td>
 <td>FST</td>
 <td>11370-31</td>
 <td></td>
 </tr>
 <tr>
 <td>Cunningham Spinal Adaptor</td>
 <td>Stoelting Co.</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Kwik-Cast sealant</td>
 <td>WPI</td>
 <td>#KWIK-CAST </td>
 <td></td>
 </tr>
 <tr>
 <td>Ventilator</td>
 <td>CWE Inc</td>
 <td>SAR-830/AP</td>
 <td></td>
 </tr>
 <tr>
 <td>Capnograph</td>
 <td>CWE Inc</td>
 <td>&mu;capstar</td>
 <td></td>
 </tr>
 <tr>
 <td>Heating blanket</td>
 <td>Harvard Apparatus</td>
 <td>507221F</td>
 <td></td>
 </tr>
 <tr>
 <td>Intracellular amplifier</td>
 <td>Axon Instruments</td>
 <td>Axoclamp 2B</td>
 <td></td>
 </tr>
 <tr>
 <td>Pipette puller</td>
 <td>Sutter Instruments</td>
 <td>P-97</td>
 <td></td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>Sigma-Aldrich</td>
 <td>P9333-500G</td>
 <td></td>
 </tr>
 </tbody>
</table>

simultaneous intracellular recording of a lumbar motoneuron and the force produced by its motor unit in the adult mouse in vivo

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 vivo2-4 , including our team who published a new preparation which allowed us to obtain very stable recordings of motoneurons in vivo in adult mice5,6. 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 circuitry7-9, or reproducting human disease10,11, we expect this technique to be an essential tool for the study of spinal motor system.

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...

Watch this Scientific Journal Video about Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo at JoVE.com

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

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.

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

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 ...

simultaneous-intracellular-recording-lumbar-motoneuron-force-produced

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JoVE Journal

Neuroscience

14.6K Views.  Northwestern University Feinberg School of Medicine. 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.

Video: Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse 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 ...

A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

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 ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

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 ...

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 ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.