Intracellular Whole-Cell Patch-Clamp Recordings of CorticalNeurons in Awake Head-Restrained Mice

Sylvain Crochet

Abstract

Membrane potential dynamics resulting from the integration of thousands of synaptic inputs and intrinsicmembrane properties underlie the generation of action potential in neurons of the central nervous system.The investigation of membrane potential dynamics is, therefore, of major importance to the understandingof brain function. This level of neuronal activity can only be assessed by measuring differences of potentialbetween the inside and the outside of a neuron, i.e., intracellular recording. In mammals, this approach hasbeen so far mainly restricted to reduced preparations in vitro and more recently to the intact brain inanesthetized animals. Such preparations do not reproduce the complexity and the diversity of the brainactivities observed in behaving animals and are, therefore, of limited interest to the understanding ofcomplex brain processing and cognitive functions. Recently, we have developed an approach that enablesintracellular recordings of cortical neurons in awake behaving mice. The mechanical stability of the brainbeing the main technical issue, it has been successfully overcome by (1) using “blind” whole-cell patch-clamp technique conferring higher stability in the initial phase of the recording, (2) implanting mice withlight metal posts that enable painless and stable fixation of the head, (3) habituating the animal to avoidlarge and brisk body movements during the recording session, and (4) reducing the size of the craniotomyto minimal to prevent large brain pulsations and edema. This technique has been successfully applied to theinvestigation of cortical sensory processing during active sensing in the mouse whisker system and has beenexpanded to simultaneous double intracellular recordings or combined with other recording techniques,such as local field potentials or two-photon microscopy.

Key words: Patch clamp, Intracellular recording, In vivo, Cortex, Behaving animal

1. Introduction

Information is encoded in the central nervous system by the activegeneration of action potentials (APs) in neurons, which areinterconnected through chemical synapses. When an AP is generatedin one neuron, it propagates along its axon to the terminals,where it releases neurotransmitter into the synaptic cleft. The neuro-transmitter binds with its receptors on postsynaptic neurons, openingion channels that generate unitary postsynaptic potentials (uPSPs).Depending upon which neurotransmitter is released, these uPSPscan be excitatory or inhibitory in nature. Due to the largely

Neuromethods (2012) 67: 219–235DOI 10.1007/7657_2011_7© Springer Science+Business Media, LLC 2011Published online: 25 November 2011

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interconnected nature of most neuronal networks in the centralnervous system, neurons have to integrate uPSPs from thousandsof presynaptic neurons (1, 2). The integration of excitatory andinhibitory inputs is done across space (location of the synapse onthe dendritic arbor) and time (summation, short-term plasticity)resulting in membrane potential (Vm) subthreshold fluctuations(3–6).When these fluctuations bring theVmclose to spike threshold,voltage-gated sodium channels open and generate an AP thatimpact several other postsynaptic neurons.

Recording APs with extracellular electrodes in intact animalshas been the primary way of assessing neuronal code and brainfunctions since the very beginning of electrophysiology by corre-lating neuronal discharge with sensory stimuli or animal behavior(7–9). Indeed, APs are high-amplitude (40–60 mV) voltage deflec-tions that can be easily recorded a few micrometers away fromthe cell using even relatively big metal electrodes (tens of mm indiameter). Extracellular recordings are, thus, less sensitive to move-ments between the electrode and the recorded neuron and canbe performed in awake head-restrained (10–13) or even freelymoving behaving animals (14–16). More recently, the paralleldevelopment of electrodes allowing for better spike detection/sorting and multisite recordings (silicon probes), as well as high-capacity multichannel acquisition systems, enabled simultaneousrecording of large neuronal population in freely moving behavinganimals (17–20).

If recording neuronal discharge is of critical importance tothe understanding of brain functions, the synaptic mechanismsleading a neuron to fire an AP are also of equal importance tounderstand how the neuronal code is elaborated. However, therecording of subthreshold Vm fluctuations is of much higherdifficulty because it can be achieved only by recording the differ-ence of potential between the inside and the outside of a neuron,which requires a direct access of the tip of the electrode to theinside of the neuron (intracellular recording). To do so, severaltechnical difficulties must be overcome. First, the tip of the elec-trode must be smaller than the soma of the neuron (for instance,the soma of a cortical neuron in the mouse is about 10–20 mm indiameter). Second, accessing the inside of a neuron implies break-ing the integrity of the cellular membrane, which may cause leaksbetween the intracellular and extracellular spaces resulting in thedepolarization of the cell and ultimately its death. It is essential,then, to establish a seal between the membrane of the recorded celland the electrode to prevent any leak. Finally, because of thesmall size of the cell, any small movement between the electrodeand the tissue may result in artifacts in the recorded Vm or inthe electrode going out or through the cell.

Two approaches have been developed in parallel over the lastdecades to perform intracellular recordings: sharp electrode

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recordings and whole-cell patch-clamp recordings. Sharp electro-des are thin glass electrodes with very small tip and high resistance(20–80 MO). The tip of the electrode has to physically penetratethe cell to access intracellular space (Fig. 1a). The penetration ofthe electrode produces a leak and a depolarization of the neuron,which is compensated by the injection of hyperpolarizing currentthrough the pipette. Over time and in the absence of movementbetween the electrode and the cell, something similar to a sealforms and a stable recording can be obtained without currentinjection (Fig. 1a). For patch-clamp technique, glass electrodes

Fig. 1. Evolution of intracellular recordings over time. (a) Schematic representation of the evolution of sharp electrodeintracellular recording. In 1, the tip of the glass pipette contacts the cell membrane. In 2, the pipette has been advancedfurther to penetrate the cell membrane, which results in leaky membrane and cell depolarization. This is compensated byinjection of hyperpolarizing current through the pipette. In 3, over time and in the absence of movement between the cell andthe electrode, a seal forms around the tip of the pipette and hyperpolarizing current can be removed. In this configuration,the recording is stable and sustains moderated tissue movements. (b) Schematic representation of the evolution of whole-cell patch-clamp recording. In 1, the tip of the glass pipette contacts the cell membrane and a gigaseal is established bygentle suction. In 2, a short negative pressure pulse applied to the pipette breaks the membrane to establish the whole-cellconfiguration. In this condition, thanks to the gigaseal and the elasticity of the cell membrane, the recording is stable andsustains moderated movements between the cell and the electrode. In 3, over time, whole-cell recordings tend todeteriorate due to increased access resistance (Rs) or the dialysis of the intracellular space.

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are bigger, with lower resistance (4–6 MO) and smooth tip. Thetip is approached against the cell membrane and a small suction isapplied to the pipette to establish a seal of very high resistance(gigaseal) before opening the membrane to access intracellularspace (Fig. 1b). The elasticity of the cell membrane attached tothe pipette by the gigaseal renders the patch-clamp approachmuch less sensitive to tissue movements than the sharp electrodeapproach in the early stage of the recording (Fig. 1). The main issuewith the patch-clamp approach is that in whole-cell configurationthe important exchanges between the intracellular milieu and thepipette solution may wash out secondary messengers and thereforeaffect the physiology of the cell (intrinsic properties, establishmentof plasticity, . . .) (Fig. 1b). Another issue with patch-clamp record-ing is that the access resistance tends to deteriorate more rapidlydue to the closure of the membrane. As a result, high-frequencycomponents of the signal, such as APs, can be attenuated (low-passfilter) (Fig. 1b).

Due to different technical issues (mechanical stability, possibilityto approach the neuron with the electrode under visual control),intracellular recordings have long been restricted to in vitropreparations (isolated cells, culture of neurons, or slices of braintissue) before being adapted to intact anesthetized animals. Thetwo main issues have been the stabilization of the preparation toprevent movements of the brain and the development of “blind”recording techniques. The stabilization can be achieved easily inanesthetized animal by different procedures, like muscular paralysisusing curare, opening of the cisterna magna to prevent brain pulsa-tion, pneumothorax and hip suspension, and covering the craniot-omy with agarose. The blind recording technique consists inadvancing the recording electrode by small steps into the braintissue until a contact with a neuron can be detected. Both sharpelectrodes and whole-cell patch-clamp techniques have been usedin vivo with their own advantages and disadvantages. Intracellularsharp recordings are more sensitive to tissue movement during thephase of stabilization, but once stabilized last longer than whole-cell patch-clamp recordings on average (up to several hours) withmore stable access resistance and less dialysis of the intracellularspace (Fig. 1). Only very recently, intracellular recordings techniquehas been expanded to nonanesthetized animals during naturalsleep–wake cycle or quiet wakefulness with either sharp electrodes(21–23) or patch-clamp technique (23, 24). But the applicationof intracellular recordings techniques to higher brain functions,such as active sensory perception, has not been possible so fardue to the difficulty of performing intracellular recordings inawake behaving animals. However, studying sensory perceptionin awake behaving animals is of major interest because neuronaldynamics and sensory processing are dramatically affected by theanimal’s behavior (21, 25–28).

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We describe here a methodological approach that we havedeveloped allowing for intracellular recordings in awake behavingmice. We detail how the major difficulty, i.e., the movementsbetween the cell and the electrode, can be overcome. We haveachieved optimal stability of the recording thanks to the use of“blind” patch-clamp technique, which is less sensitive to move-ments than sharp electrodes; implantation of a light metal post onthe head of the animal, enabling painless and stable head fixation(Fig. 2); a period of habituation of the mouse to the procedure ofhead fixation to avoid large and brisk body movement during therecording session; and reduction of the size of the craniotomy.

Fig. 2. Head post and holder for intracellular recording in awake head-restrained mice. (a) Pictures of a metal post to beimplanted on the skull of a mouse (scale bar, 10 mm). (b) Pictures of the head-post holder. Note the two orthogonal rotationaxes allowing the adjustment of the position of the mouse head. The head post is fixed thanks to a nut. (c) View of theintracellular setup configuration with two microdrives allowing for simultaneous positioning of two electrodes (local fieldpotential + intracellular; juxtacellular+intracellular; dual whole-cell recordings).

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Assessing Vm dynamics in awake behaving animals providesnew insights into brain function, opening the possibility ofdissecting the synaptic mechanisms that drive neuronal networks.We have applied this technique to the somatosensory system ofthe mouse, which constitutes a very attractive model of activesensing. Mice and rats have a very-well-organized system ofvibrissae (or whiskers) with a topological organization conservedfrom the whisker pad to the primary sensory cortex, with eachsingle-whisker activating neuron preferentially within a singlebarrel column (29, 30). Mice and rats use their whiskers to activelyexplore their environment by moving their whiskers back and forthin a rhythmic manner (whisking) and are able to perform very finediscrimination of texture or object position (14, 15, 31–34).We have combined intracellular recordings with high-speed videofilming of whisker movements to correlate Vm dynamics withwhisker behavior and active sensing (35). This technique has beenfurther expanded in our laboratory to simultaneous dual whole-cell recordings and targeted whole-cell recordings of GABAergicinterneurons using two-photon microscopy in awake mice, show-ing the impact of behavior on neuronal discharge and correlation ofVm fluctuations between pyramidal and GABAergic interneuronsin the primary somatosensory cortex (25, 36). Other laboratorieshave also recently successfully implemented a very similar approachto perform whole-cell patch-clamp recordings in awake head-restrained rodents in the neocortex or hippocampus (23, 37).

The major weakness of the present approach are as follows:(1) We have noticed that 2–3 weeks after the implantation, the duramater underneath the exposed bone in the recording chambertends to grow thicker and becomes more vascularized—openinga clean craniotomy in this condition becomes much more difficultand the success rate of the patch-clamp recording is significantlyreduced. (2) The behavioral tasks that can be used are obviouslylimited by the reduced motor outputs and the increased diffi-culty of training an animal, especially a mouse, while head fixed.But it should be noted that despite recent successful attempts, thesuccess rate of intracellular recordings in freely moving animalsso far is very low (about 15%) (38, 39). In addition, several labora-tories have recently developed more sophisticated behavioral tasksin head-restrained rodents (34, 37). The use of intracellularwhole-cell recording in awake head-restrained rodents is, therefore,a promising technique, which will certainly develop in the future,in combination with other electrophysiological or imagingapproaches.

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2. Materials

2.1. Experimental

Animals of InterestOur experiments have been conducted on 4–8-weeks-old C57Bl6Jmice from Janvier (see Note 1). All experiments were carried outin accordance with the Swiss Federal Veterinary Office. Make surethat the described protocol conforms to institutional and nationalregulation.

2.2. Implantation l Vapomatic anesthetic vaporizer (VIP3000 Isoflurane, MDSMatrx) and Attane Isoflurane (Provet).

l Stereotaxic device: Nose clamp (custom made) or small animalstereotaxic frame (Kopf).

l Heating blanket: DC Temperature Control Module, heatingpad, and immersion heating rod (FHC).

l Surgical instruments: Fine scissors, disposable scalpel bladeswith handle, spatula.

l Head post (custom made): We have developed an L-shapedhead post that enables maximum contact surface with the skulland good access to most of brain regions (Fig. 2a). The longpart of the head post is positioned contralaterally to the regionof interest and the short part over the cerebellum; the shape canbe modified for particular applications (bilateral recordings,access to the cerebellum). An angle of ~140� between thepart in contact with the skull and the part fixed to the holder(Fig. 2b) provides a better access to the craniotomy. The partfixed to the holder has a trapezoidal shape that ensures a goodstability between the head post and the holder.

l Ophthalmic gel (Viscotears, Novartis).

l Topical antiseptic (Betadine).

l Hydrogen peroxide solution (2%).

l Cotton swabs.

l Cyanoacrylate glue (Loctite Super Attak 401, Henkel).

l Acrylic dental cement (Paladur pink powder 67407963 andPaladur liquid, Kaladent).

l Silicone sealant (Kwik-Cast, WPI).

l Carprofene (Pfizer).

2.3. Habituation l Head-post holder (custom made): The head-post holder hasbeen developed to provide maximum stability and flexibilitywith two orthogonal axes of rotation allowing for the adjust-ment of the position of the mouse head (Fig. 2b).

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2.4. Craniotomy l Ringer’s solution containing (in mM): 135 NaCl, 5 KCl, 5HEPES, 1 MgCl2, and 1.8 CaCl2 (adjusted to pH 7.3with NaOH).

l Stereomicroscope (MZ95, Leica) with long working distanceobjective (plan �1.0, 112 mm), magnification from 6.3 to 60with 10�/21B eyepieces.

l Lightweight drill (Model EXL-M40, Osada) and small diameterdrill bit (Komet Dental Drill Bit H1.204.005, CondorDental).

l Surgical instruments: Fine forceps (Dumont #5) and needles(U-100 Insulin, 30 G � 8 mm, BD Micro-Fine).

2.5. Recording l Glass capillaries: Bo-glass capillaries, ends fire-polished 75 mmlength OD ¼ 2.0, ID ¼ 1.4 mm (Hilgenberg).

l Pipette puller: P-97 Flaming/Brown Micropipette Puller(Sutter Instrument).

l Intracellular solution containing (in mM): 135 potassium gluco-nate, 4 KCl, 10HEPES, 10 sodium phosphocreatine, 4MgATP,0.3 Na3GTP (adjusted to pH 7.3 with KOH and to osmolarity285 mOsm/L with distilled water). The intracellular solutioncontains ATP and GTP that degrade rapidly at room tempera-ture. Stock solution must be prepared on ice and a 1-mL aliquotcan be stored at�20�C for a fewmonths. Just before the record-ing session, unfreeze the aliquot and add 3 mg/mL biocytin.Then, keep the solution on ice.

l Microdrive: Mini 4 axes, 4MRE/4MLE (Luigs & Neumann).

l Pipette pressure control:Manual seal sucker (SigmannElektronik).

l Intracellular amplifier: Multiclamp 700 amplifier (Axon Instru-ments).

l Recording system: Signals are filtered online at 10 kHz(bessel filter, Multiclamp) and digitized at 20 kHz by ITC-18(Instrutech Corporation) under the control of IgorPro (Wave-metrics).

l High speed camera (MotionPro, Redlake), synchronized withelectrophysiology through TTL pulse.

l Reference electrode: Ag/AgCl Electrode 2.0 mm diameter(WPI).

l Multichannel digital oscilloscope: TDS 2014 (Tektronic).

3. Methods

3.1. Implantation Timing: 30–45 min per animal

1. Anesthetize the animal with isoflurane (3%) in oxygen. Oncedeeply anesthetized (deep and slow breathing), the mouse

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remains unconscious for tens of seconds out of the inductionbox (see Note 2) and its head can be fixed into the nose clampor the stereotaxic frame.

2. Place the animal onto the heating blanket and insert the rectalprobe to maintain the body temperature at ~37�C. Decreaseisoflurane concentration to 2%. Apply ophthalmic gel on theeyes to prevent ocular dryness.

3. Expose the skull by removing the skin from the cerebellum tothe olfactory bulb.

4. To ensure a good anchoring of the head post, it is essential tocarefully clean the skull. First, gently clean the exposed skulland surrounded cut skin with betadine applied with cottonswabs. Retract the membranes covering the skull. Then, cleanthe skull with peroxide solution. Dry the skull with cottonswabs. Use the edge of a scalpel blade to clean the wholesurface of the skull free of periosteum. Finally, make shallowgrooves every 0.5–1 mm on the surface of the skull with thetip of the scalpel blade, except where the recording chamber isgoing to be (see Note 3).

5. Cover the whole exposed skull with a thin layer of cyanoacry-late glue. After a few minutes, the L-shaped head post is gluedonto the skull with the long bar in contact with the skullpositioned contralaterally to the region of interest (the primarysensory cortex in our case) and the short part lying over thecerebellum (see Note 4).

6. After a few minutes, the head post is securely attached to theskull with acrylic dental cement, which is also used to buildthe recording chamber. Liquid dental cement is first applied tothe whole surface of the skull (except where the recordingchamber is going to be positioned) as well as over the parts ofthe head post in contact with the skull. Prepare dental cementinto a consistency comparable with toothpaste and use it tobuild the walls of the recording chamber. The recording cham-ber is typically 5–8 mm in diameter and the chamber walls2–3 mm high (see Note 5). Once the dental cement is solid,fill the recording chamber with silicone sealant.

7. Administrate a pain killer to the mouse (carprofene, i.p.).

8. Put the mouse in a clean cage and let it recover from surgeryfor 2–3 days. Control body weight right after and in the nextdays following implantation. The animal should not lose morethan 10% of its body weight and then recover.

3.2. Habituation Timing: 3–5 days per animal

1. Put the animal onto the platform and fix the head post to theholder. The body of the animal can be slightly restrained by atub or cardboard walls (see Note 6).

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2. At the end of each session, reward the animal with a few dropsof sweet water, then release it, and put it back into its cage.

3. Control body weight during the habituation period. Theanimal should not lose weight.

4. The duration of each session should increase gradually, startingfrom 10 to 20 min up to 1–1.5 h. The number and the incre-ment in time between each session should be adapted for eachanimal depending on its behavior and the experimental protocol(see Note 7).

3.3. Craniotomy Timing: 30–40 min per animalThe craniotomy is a very critical step for the patch recording,especially when recording from superficial regions of the neocortex.The dura must be removed without bleeding in order to ensure arecording of good quality. Any trace of membrane or blood mightprevent the establishment of the gigaseal. In mice, there is very littlespace between the pia and the dura mater. Opening the durawithout damaging the superficial layer of the cortex is a delicateoperation that requires some practice. The size of the craniotomy isalso very important: the smaller it is, the more stable the recordingis (see Note 8). This procedure must be conducted using a high-magnification stereomicroscope and a good illumination of thepreparation.

1. Anesthetize the animal (see step 1, section “Implantation”).

2. Fixe the head post to the holder firmly and decrease isofluraneconcentration to 2%.

3. Remove the silicone sealant from the recording chamber andclean it with warm (~37�C) Ringer’s solution.

4. Determine the location of the craniotomy (see Note 9).

5. Drill circularly around the region of interest to thin the bone.Start with circles of ~1.5 mm centered on targeted point andprogressively reduce diameter as the bone gets thinner(see Note 10). Stop drilling before reaching the dura. Thebone should be thinned on a region of 0.2–0.4 mm in diameter(Fig. 3).

6. Insert a fine needle between the thinned bone and the dura toopen the craniotomy (Fig. 3).

7. Carefully clean if any bleeding occurs.

8. Incise the dura mater with the tip of a clean and new needle(Fig. 3). The opening of the dura can be seen by a leak ofcerebrospinal fluid (see Note 11).

9. Fill the recording chamber with Ringer’s solution and seal thechamber with silicone sealant.

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10. Put the animal in a separate cage for recovery (2–24 h) until therecording session.

3.4. Recording Timing: 1–3 h per animal

1. Pull the pipettes (long tip, 4–7 MO).

2. Unfreeze one aliquot of intracellular solution and add thebiocytin.

3. Place the animal on the recording setup and fix the head post tothe holder.

4. Remove the silicone sealant and gently clean the craniotomy(see Note 12).

5. Position the reference electrode in or at the border of therecording chamber. A small piece of plasticine can be used tomaintain the reference electrode in position.

6. Fill the patch pipette with fresh intracellular solution.

7. Insert the pipette in the pipette holder and apply positivepressure to the pipette (100–150 mb).

Fig. 3. Procedure for the craniotomy. Schematic representation of the procedure to open acraniotomy for cortical intracellular recording. The bone is thinned by drilling circularlyand progressively reducing circle diameter. The bone is then open by inserting a thinsharp needle between the bone and the dura. After gentle cleaning to remove possibleblood trace, a sharp needle is used to open the dura.

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8. Under stereomicroscope guidance, position the tip of thepipette into the craniotomy, on the surface of the brain.

9. The blind patch technique consists in advancing the patchpipette by steps of 2 mm into the brain until the tip of thepipette contacts a cell, which can be seen as a sudden increasein the resistance of the patch pipette. This procedure is con-ducted in voltage-clamp mode by constantly applying 10-mVsquare test pulses at 20–50 Hz and monitoring the currentdeflection recorded by the patch pipette. A decrease in thedeflection indicates an increase in the pipette resistance.

10. Using the wheel of the control pad, slowly advance the pipetteinto the brain by ~50 mm, then release the pressure to50–70 mb, and check the resistance of the pipette (see Note13). Advance the pipette by a few 2-mm steps. If the resistanceof the pipette does not increase, the pipette can be slowlyadvanced to the region of interest using the wheel of thecontrol pad. Constantly check the resistance of the pipette;the resistance may increase due to contact with a cell or ablood vessel but generally returns to baseline after tens of mm.If the resistance remains high, withdraw the pipette a bit andapply higher positive pressure until the resistance returns tobaseline (see Note 14).

11. Once the recording site is reached, decrease the pressure to20–30 mb and advance the pipette by 2-mm steps, until acontact occurs. A good contact appears as an abrupt approxi-mately twofold increase in input resistance (Fig. 4a). Releasethe positive pressure. Immediately and gently apply mild neg-ative pressure (suction) while simultaneously and progres-sively decreasing the holding potential to �70 mV, until agigaseal is formed (Fig. 4a) (see Note 15). At this stage, theelectrode capacitance can be compensated. Then, open themembrane by applying short and transient negative-pressurepulses to the pipette until the membrane breaks (Fig. 4a).Then, switch the amplifier to current-clamp mode (Fig. 4b).

12. The good quality of the recording is attested by mean Vm< �40 mV, overshooting APs with amplitude >40 mV.Low-amplitude, broad APs generally result from high accessresistance, which may indicate that the membrane is notopened enough. Applying slight negative or positive pressureas well as moving the pipette one step back may correct forthe bad access resistance, but this may also cause the loss ofthe recording.

13. Once a whole-cell recording of good quality is obtained, pro-ceed with your protocol. An example of correlation betweenVm dynamics from a layer 2/3 pyramidal neuron in the barrelcortex and whisker behavior is depicted in Fig. 4b–e.

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Fig. 4. Blind whole-cell patch-clamp recording in an awake behaving mouse. (a) Schematic representation of thesuccessive phases leading to whole-cell configuration. In voltage clamp mode, 10-mV test pulses are applied to thepipette at 20–50 Hz while monitoring the current response on the oscilloscope. A constant 15–30-mb positive pressure isapplied to the pipette, which is advanced in 2-mm steps into the brain. A contact with a cell is detected as a suddenincrease in the pipette resistance (decrease in current in response to test pulse). The pressure is then released producing afurther increase in the pipette resistance. A gentle suction (negative pressure) is applied to the pipette while the holdingpotential is progressively decreased to�70 mV, until a gigaseal is formed. After capacitance and transient compensation,short, negative-pressure pulses are applied to the pipette to break the cell membrane and establish the whole-cellrecording. The amplifier can then be switched to current-clamp mode. (b) Schematic representation of the recordingconfiguration in awake head-restrained mouse and an example of a whole-cell patch-clamp recording of a layer 2/3pyramidal cell within C2 barrel column during a sequence of active touches. Top trace, whisker angular position (WP);bottom trace, membrane potential (Vm); gray bars indicate contacts between the whisker and the object. (c) Reconstructeddendrites and descending axon within barrel map (normal view). The recorded neuron was filled with biocytin and stainedwith an ABC kit and diaminobenzidine for post hoc anatomical identification and localization relative to barrel map.(d) Simultaneous high-speed filming of the whisker behavior allowed for identification of whisker–object contact time.(e) Averaged WP and Vm triggered by the onset of active contacts. (e) Averaged WP and Vm triggered by the onset of theactive contacts (black line).

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14. At the end of the recording, the pipette is slowly withdrawn andthe animal can be Transcardially perfused with PFA solution forpost hoc anatomical identification of the recorded neuron(Fig. 4c). To increase the chance of recovery of the cell anat-omy, the pipette should preferentially be withdrawn while therecording is still of good quality.

4. Notes

1. A similar approach can be used with rats. However, rats aremuch stronger than mice and can easily break their implantsduring the habituation procedure. It is, thus, recommended touse anchoring screws during the implantation and to processvery gently and progressively during the habituation (23, 40).

2. For short implantations (up to 45 nm), gas anesthesia can besubstituted by a mixture of Ketamine and Xylazine adminis-tered by i.p.

3. The skull of the mouse is very soft, so particular attentionshould be paid not to apply excessive pressure with the cottonswabs or the scalpel blade.

4. The head post can be stereotaxically positioned using a mod-ified arm mount micromanipulator (Kopf) to enable reproduc-ible implantation.

5. The size of the recording chambermay be adapted to experimen-tal needs. The critical point is to position and build the recordingchamber in a way that enables a good access to the craniotomy,i.e., the needle should reach the craniotomy with an angle nothigher than 40� to the surface of the brain (Fig. 3).

6. The use of any plastic materials should be avoided in closeproximity of the animal. For example, if the animal is sittingon a plastic board, we observed very large electrical artifacts dueto static charge when the animal is moving or whisking.

7. The habituation process can be largely adapted depending on theexperimental protocol. Three to five sessions of habituation areusually necessary to perform recordings in an awake mouse.Recordings across different sleep–wake stages (wakefulness,slow-wave sleep, and REM sleep) are also possible but requireprolonged habituation (2–3 weeks) with longer sessions (3–5 h)(41). It is also possible to train an implanted animal to performsimple tasks under head-restrained condition during the periodthat precedes the recording (34).However, there are two limitingfactors to a long habituation process or training procedure:(1) when the animals are implanted for too long (more than3 weeks), the dura underneath the exposed skull grows thicker

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making the craniotomymore difficult; (2) it is also more difficult,though not impossible, to perform patch-clamp recordings inmice older than 6 weeks and our protocol for implantationmight not be appropriate for mice younger than 4 weeks. There-fore, it is not recommended towaitmore than2–3weeks betweenthe implantation and the recording session.

8. For combination with two-photon microscopy, the size of thecraniotomy must be larger (1–2 mm in diameter). To preventtissue movements and edema, the brain is stabilized by coveringthe craniotomy with agarose and a thin glass coverslip.

9. The location of the craniotomy can be determined in differentways. Stereotaxic coordinates can be used; in this case, a stereo-taxic frame must be used during the implantation and thetargeted location can be marked during the implantation.A major advantage of the mouse is that the skull, whenhydrated, is transparent enough to perform intrinsic imagingwithout opening or thinning. For experimenters interested insensory perception, the exact location of the craniotomy canthus be functionally determined using intrinsic optical imaging(28, 35). In any case, it is recommended to choose a locationavoiding big blood vessels.

10. The skull of the mouse is very thin and transparent whenhydrated but becomes opaque when dry. It is easier to drillwhen the bone is dry, but it is recommended to regularlycontrol the progression of the drilling by filling the chamberwith Ringer’s solution.

11. Alternatively, the opening of the dura can be done at thebeginning of the recording session, just prior to the insertionof the pipette. This is actually recommended if the recordingsession occurs the day after the craniotomy.

12. Depending on the duration of the recovery period after thecraniotomy, it might be necessary to reopen the dura with a fineneedle.

13. If the resistance of the pipette is high after its insertion into thetissue or progressively increases when advancing it, it meansthat the tip of the pipette is obstructed probably because thecraniotomy was not perfectly clean or the dura not completelyremoved. One can try to clean the tip of the pipette by increas-ing the positive pressure applied to the pipette and advancingthe pipette by 50–100 mm. If the pipette cannot be cleaned,it must be changed and it is recommended to clean the crani-otomy again.

14. The pressure used to advance the pipette into the tissue mightbe adjusted depending on which structure is targeted; reachingdeep structures may require passing through areas rich in fibersor even through ventricles with high risk of obstructing thetip of the pipette.

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15. Sometime, the establishment of a gigaseal is difficult despitethe application of negative pressure to the pipette. Depolariz-ing the holding Vm to �80/�90 mV may help forming theseal. The holding Vm should then be set back to �70 mVbefore breaking the membrane to establish the whole-cellconfiguration.

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