Supported by NSF grant 0920060, NIH grant DC004395, NHMRC grant 1009482, NSW Office of Science and Medical Research, and the Garnett Passe and Rodney Williams Memorial Foundation.

1. Head-restraint Overview
<ol>
	<li>To assemble a custom-built head-post, insert a 1/16" stainless steel roll-pin perpendicularly into a hole drilled in a 3/32" stainless steel rod to form a cross. The vertical piece of the head-post should be about 20 mm and the horizontal cross piece about 15 mm. Tap one end of the vertical rod to accept a #1-72 screw. (Fig. 1).</li>
	<li>During surgery and recording, the head-post is secured in a custom-built brass or aluminum mounting bar with a #1-72 screw (Fig. 2). The long piece of the mounting bar should be about 90 mm long and 5-10 mm wide. The head-post fits into a groove cut into a 15 mm extension off the long bar that is angled at 45 deg. To prevent head rotation, the crosspiece of the head-post nestles into a small groove on the bottom of the mounting bar.</li>
	<li>The mounting bar in then attached to a custom-made aluminum mounting block (Fig. 3). The mounting block is approximately 30 mm x 30 mm x 25 mm. This mounting block is attached to a micromanipulator so that the position of the head-post can be accurately placed along the midline and at Bregma. The mounting block is made of two pieces fitted together with screws. A cut-out in the block allows the mounting bar to slide in and be secured with the screws. The mounting bar is parallel to the table surface. During recording, secure the bar in a custom stereotaxic frame (Fig. 4). The stereotax is designed to mount the bar in the same position as in the mounting block during surgery. This arrangement ensures that the mouse will always be aligned to the stereotaxic frame across experimental sessions.</li>
	<li>Cut a tungsten rod (diameter .010") to approximately 5 mm length, with approximately 1 mm bent at 45&deg;. This rod serves as the ground pin (Fig. 1).</li>
</ol>
2. Stereotaxic Alignment for Craniotomy
Note: All procedures outlined below follow standard aseptic surgical techniques and have been approved by the Washington State University Animal Care and Use Committee and the Animal Ethics Committee of the Garvan Institute.
<ol>
	<li>Anesthetize the mouse by placing in an induction chamber with 5% isoflurane. The lack of a reaction to tail and/or toe pinch ensures that the animal is fully anesthetized.</li>
	<li>Place the mouse in a rodent stereotaxic frame outfitted with a mouse adapter (Stoelting). To orient the mouse correctly, place the teeth inside the hole of the bite bar and tighten the nose clamp so it is snug. Place the mouse as straight as possible in the bite bar and nose clamp. Place a face mask made from a piece of latex glove over the mouse's nose, and keep the isoflurane at 5% until the mouse's respiration rate is approximately 1 breath/sec. Turn the isoflurane to 1.5 - 2.0%. Secure the head using the ear-bars being careful to not puncture the ear drum. Bars should be snug but not penetrating too deeply into the ear canals.</li>
	<li>Place a heating pad underneath the mouse.</li>
	<li>Apply ophthalmic ointment to prevent the eyes from drying out.</li>
	<li>Shave the scalp from between the ears up to the eyes with a small shaver or fine-point scissors.</li>
	<li>Clean and sterilize the scalp by wiping with chlor-hexidine (or betadine) scrub followed by alcohol rinse repeated 3 times.</li>
	<li>To remove the skin over the top of the skull make an incision along the midline from the back of the head to the eyes.</li>
	<li>Scrape the periosteum to the edges of the incision with a scalpel. If required (depending on experimental target), carefully retract the musculature at the back of the neck with forceps, allowing the muscle to tear along fascicles to minimize bleeding. Apply gelfoam underneath scalp tissue to minimize moisture and prevent the skin from moving over the skull.</li>
	<li>Dry the skull surface with a cotton-tipped applicator moistened with isopropyl alcohol to enhance the visualization of skull landmarks.</li>
	<li>Align the skull to standard stereotaxic coordinates13. Insert a fine-point probe in the electrode holder and attach it to the stereotaxic micromanipulator. A sharpened metal rod works well for this purpose (Fig. 5).</li>
	<li>Use the point to identify Bregma (Fig. 5A) and Lambda (Fig. 5B) on the skull. Move the probe back-and-forth between these points, checking that the lateral-medial coordinate for each location differs by no more than 50 &mu;m. Reposition the head as needed by carefully loosening the ear-bars, shifting the head and ear-bars laterally, and retightening the ear-bars.</li>
	<li>Verify that the dorsal-ventral height at each location differs by no more than 50 &mu;m. Adjust the height of the nose-clamp, which will rotate the skull around the axis of the ear-bars, to level the pitch of the skull if necessary. The height of the ear-bar might also need to be adjusted.</li>
	<li>Use an atlas13 to identify the coordinates of the target of interest (Fig. 5C) relative to Bregma. Mark the skull above the target (e.g., an "X" or use India ink to dot four corners of a square) where a craniotomy will be made (Fig. 5D).</li>
</ol>
3. Head-post and Ground Pin Installation
<ol>
	<li>Use etch gel to clean skull. Use the applicator to scrub and spread etch gel over the surface (10 sec). Wash with water and dry completely. Apply the primer for dental cement to the skull surface using applicator. Spread adhesive with new applicator. Cure the adhesive with a UV light-gun (1 cycle).</li>
	<li>Choose a location for the ground pin that is distal to the target site. Carefully bore a small hole in the skull using the tip of a #65 miniblade scalpel just wide enough for the short end of the ground pin to enter and rest upon the meninges. The ground pin should be oriented so that it is pointing away from Bregma and the future head-post (Fig. 6A).</li>
	<li>Secure the head-post to the mounting bar and block and attach to the micromanipulator. By moving the micromanipulator in three dimensions, position the head-post over Bregma, just touching the skull (Fig. 6B).The base of the head-post should lie flat on the skull.</li>
	<li>Using dissection probes, apply a small amount of dental cement around the head-post and ground pin (Fig. 6C). Press the cement down to ensure good contact with the skull. Note: Do not use too much cement or it will not harden evenly. Cure the cement with a UV light-gun (3-4 cycles from multiple angles).</li>
	<li>Apply the second stage of cement and build up the base around the head-post and cover the grooves of the head-post (Fig. 6D).</li>
</ol>
4. Craniotomy
<ol>
	<li>Locate the craniotomy reference mark, which will be partially obscured by the primer. Use a #65 scalpel to mark a 3 mm x 3 mm square around this site and apply a drop of lidocaine to reduce sensitivity and to minimize bleeding.</li>
	<li>Make multiple scores on each side of the square (as many as 10), being careful not to cut through the bone and into the brain (Fig. 7A).</li>
	<li>Once the bone flap becomes loose, use the tip of the scalpel to pry it up, leaving the dura mater intact (Fig. 7B).</li>
	<li>Cover with bone wax.</li>
</ol>
5. Post-surgery
<ol>
	<li>Turn off gas anesthesia and remove the mouse from the stereotaxic frame.</li>
	<li>Use a cotton-tipped applicator to apply Lidocaine ointment around exposed skin to deaden sensation around wound. Use another cotton-tipped applicator to apply Neosporin to the exposed skin. Inject ketoprofen (5 mg/kg IM or IP based on preference of individual IACUC veterinarians) as a post-surgery analgesic. Mouse will generally be up and moving within minutes of gas cessation.</li>
	<li>Place mouse in a warm cage and monitor until it is ready to return to animal housing.</li>
	<li>The animal should be checked on at least once per day after it has recovered from surgery. Watch for signs of poor eating and/or drinking and listless behavior. If pain is suspected, provide ketoprofen every 24 hours (same dose as at the end of post-surgery) until alleviated. Lidocaine can be locally applied to the wound if the animal is scratching or shows signs of discomfort. Animals typically recover from surgery without complications or pain.</li>
</ol>
6. Recording and Dye Injection
<ol>
	<li>Wait at least one day after head-post surgery before using the mouse for recordings.</li>
	<li>Place mouse in a foam-restraining device molded to the mouse's body. The mouse has already been adapted to this device, thereby reducing its anxiety. Typical adaptation is to handle the mouse every day after weaning and for approximately one week prior to surgery and experiments, place the mouse in the foam device for 1-5 minutes. Suspend device in the stereotaxic frame (Fig. 4).</li>
	<li>Secure head-post to mounting bar, which is now fixed to the stereotaxic frame (Fig. 4).</li>
	<li>Connect to the ground pin with a wire terminated with a blunt 22-gauge hypodermic needle tip.</li>
	<li>Remove bone wax over the craniotomy. Remove dura if necessary using the tip of a miniblade scalpel.</li>
	<li>Drive electrode to proper stereotaxic location and begin recording (Fig. 8). For only electrophysiological recordings, a variety of electrodes can be used. The choice of electrode is dependent on the location and type of recordings required. The choice of electrode for the awake mouse recordings will be the same as what a researcher uses in an anesthetized mouse. For combined electrophysiological recordings with dye/tracer injections, a micropipette electrode is used10. The micropipette is filled with the dye/tracer of choice so that it can be iontophoretically injected at the end of electrophysiological experiments. Multibarrel electrodes can also be used so that pharmacological manipulations can be made. The multibarrel electrode is mounted onto a single recording micropipette (Fig. 8). These techniques are standard in the literature and are not different for anesthetized versus awake recordings.</li>
	<li>Recordings can be made for 4-5 hours over 3-4 consecutive days.</li>
	<li>At the end of the recording session, the dye/tracer can be injected. Because there are no pain receptors in the brain, the iontophoretic injection of the dye or tracer into the brain is unlikely to cause discomfort. However, once the electrophysiological response properties of the region where the injection is to be made are characterized, the animal could be anesthetized with isoflurane for the injection. This would eliminate any potential unwanted sensations. Because the dye/tracer is already in the micropipette, the active and ground leads can be switched from the electrophysiological recording set-up to the current generator to iontophoretically inject the dye/tracer. Use appropriate protocol for the dye/tracer in the electrode. Remove the animal from the headpost restraint and body restraint and return to its home cage.</li>
	<li>At the end of the experiment, follow appropriate protocol for euthanasia and tissue processing to recover injection site and labeling.</li>
</ol>
7. Representative Results
Successful installation of the head-post allows the experimenter to record single and multiunit responses from an unanesthetized, awake mouse over multiple days. The restraint system allows for stable electrophysiological recordings of single neurons in the brain. Excellent isolation of single units and strong responses to stimuli can be recorded from the same brain structure over multiple days, for example in the mouse inferior colliculus (Fig. 9A, day one; Figure 9B, day two). With a good head-post installation, each mouse will be consistently aligned to the stereotaxic apparatus and the mouse stereotaxic atlas,13 resulting in reliable localization of particular brain nuclei. This reliable localization allows for injection of dyes and tracers into specific structures after multiple recording days. For example, to assess descending projections from the auditory midbrain to the auditory brainstem, biotinylated dextran amine (visualized using diaminobenzidine as chromagen) can be injected into the mouse inferior colliculus (Fig. 10A) after electrophysiologically identifying neuronal response properties at the site of injection (in Fig. 10A, a neuron was recorded with a best frequency of 51 kHz), and after tissue processing, anterograde labeling in the contralateral dorsal cochlear nucleus can been visualized and plotted (Fig. 10B). Photomicrographs at higher resolution can also be obtained to view anterogradely labeled axons and terminals (Fig. 10C).
<img alt="Figure 1" src="/files/ftp_upload/3755/3755fig1.jpg" /> 
Figure 1. 
<img alt="Figure 2" src="/files/ftp_upload/3755/3755fig2.jpg" /> 
Figure 2. 
<img alt="Figure 3" src="/files/ftp_upload/3755/3755fig3.jpg" /> 
Figure 3. 
<img alt="Figure 4" src="/files/ftp_upload/3755/3755fig4.jpg" /> 
Figure 4. 
<img alt="Figure 5" src="/files/ftp_upload/3755/3755fig5.jpg" /> 
Figure 5. 
<img alt="Figure 6" src="/files/ftp_upload/3755/3755fig6.jpg" /> 
Figure 6. 
<img alt="Figure 7" src="/files/ftp_upload/3755/3755fig7.jpg" /> 
Figure 7. 
<img alt="Figure 8" src="/files/ftp_upload/3755/3755fig8.jpg" /> 
Figure 8. 
<img alt="Figure 9" src="/files/ftp_upload/3755/3755fig9.jpg" /> 
Figure 9. 
<img alt="Figure 10" src="/files/ftp_upload/3755/3755fig10.jpg" /> 
Figure 10.

<ol>
	<li>Evans, E. F., Nelson, P. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+responses+of+single+neurones+in+the+cochlear+nucleus+of+the+cat+as+a+function+of+their+location+and+the+anaesthetic+state.">The responses of single neurones in the cochlear nucleus of the cat as a function of their location and the anaesthetic state.</a> Exp. Brain Res. 17, 402-427 (1973).</li><li>Joris, P. X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Response+classes+in+the+dorsal+cochlear+nucleus+and+its+output+tract+in+the+chloralose-anesthetized+cat.">Response classes in the dorsal cochlear nucleus and its output tract in the chloralose-anesthetized cat.</a> J. Neurosci. 18, 3955-3966 (1998).</li><li>Populin, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthetics+change+the+excitation/inhibition+balance+that+governs+sensory+processing+in+the+cat+superior+colliculus.">Anesthetics change the excitation/inhibition balance that governs sensory processing in the cat superior colliculus.</a> J. Neurosci. 25, 5903-5914 (2005).</li><li>Young, E. D., Brownell, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Responses+to+tones+and+noise+of+single+cells+in+dorsal+cochlear+nucleus+of+unanesthetized+cats.">Responses to tones and noise of single cells in dorsal cochlear nucleus of unanesthetized cats.</a> J. Neurophysiol. 39, 282-300 (1976).</li><li>Donoghue, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contrasting+properties+of+neurons+in+two+parts+of+the+primary+motor+cortex+of+the+awake+rat.">Contrasting properties of neurons in two parts of the primary motor cortex of the awake rat.</a> Brain Res. , 333-3173 (1985).</li><li>Westby, G. W., Wang, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+floating+microwire+technique+for+multichannel+chronic+neural+recording+and+stimulation+in+the+awake+freely+moving+rat.">A floating microwire technique for multichannel chronic neural recording and stimulation in the awake freely moving rat.</a> J. Neurosci. Methods. 76, 123-133 (1997).</li><li>Suga, N., O'Neill, W. E., Manabe, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cortical+neurons+sensitive+to+combinations+of+information-bearing+elements+of+biosonar+signals+in+the+mustache+bat.">Cortical neurons sensitive to combinations of information-bearing elements of biosonar signals in the mustache bat.</a> Science. 200, 778-781 (1978).</li><li>O'Neill, W. E., Suga, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Target+range-sensitive+neurons+in+the+auditory+cortex+of+the+mustache+bat.">Target range-sensitive neurons in the auditory cortex of the mustache bat.</a> Science. 203, 69-73 (1979).</li><li>Suga, N., O'Neill, W. E., Manabe, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Harmonic-sensitive+neurons+in+the+auditory+cortex+of+the+mustache+bat.">Harmonic-sensitive neurons in the auditory cortex of the mustache bat.</a> Science. 203, 270-274 (1979).</li><li>Portfors, C. V., Felix, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=2nd+Spectral+integration+in+the+inferior+colliculus+of+the+CBA/CaJ+mouse.">2nd Spectral integration in the inferior colliculus of the CBA/CaJ mouse.</a> Neuroscience. , 136-1159 (2005).</li><li>Felix, R. A., Portfors, C. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Excitatory,+inhibitory+and+facilitatory+frequency+response+areas+in+the+inferior+colliculus+of+hearing+impaired+mice.">Excitatory, inhibitory and facilitatory frequency response areas in the inferior colliculus of hearing impaired mice.</a> Hear Res. 228, 212-229 (2007).</li><li>Portfors, C. V., Jonson, K. G., Roberts, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Over-representation+of+species-specific+vocalizations+in+the+awake+mouse+inferior+colliculus.">Over-representation of species-specific vocalizations in the awake mouse inferior colliculus.</a> Neuroscience. 162, 486-500 (2009).</li><li>Franklin, K. B. J., Paxinos, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Mouse Brain in Stereotaxic Coordinates. , (2007).</li></ol>

We have nothing to disclose.

The advantage of the head-post restraint system for electrophysiological and neuroanatomical experiments in mouse is that experiments can be conducted with unanesthetized, awake mice, eliminating potential response contamination due to anesthetic drugs. In addition, the set up can be used over multiple days to allow for more efficient animal use. 
Most components for the head-post are available off-the-shelf. Only the head-post and mounting apparatus are custom-made, both of which are relativ...

<table border="1">
 <tbody>
 <tr>
 <td>Name</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>Etch gel for cleaning skull prior to dental cement</td>
 <td>Henry Schein</td>
 <td>101-5396</td>
 <td>12/pk with 1.2 mL syringes</td>
 </tr>
 <tr>
 <td>Primer for dental cement</td>
 <td>Kerr Optibond</td>
 <td>25881</td>
 <td>8 mL bottle</td>
 </tr>
 <tr>
 <td>Adhesive for dental cement</td>
 <td>Kerr Optibond</td>
 <td>25882</td>
 <td>8 mL bottle</td>
 </tr>
 <tr>
 <td>Applicators for dental cement</td>
 <td>Kerr Optibond</td>
 <td>24680</td>
 <td>200/pk</td>
 </tr>
 <tr>
 <td>Dental Cement</td>
 <td>Charisma, Heraeus Kulzer</td>
 <td>66000085</td>
 <td>4gm Syringe Refill</td>
 </tr>
 <tr>
 <td>Tungsten Rod (Ground Pin)</td>
 <td>A-M Systems</td>
 <td>717200</td>
 <td>0.010" diameter</td>
 </tr>
 <tr>
 <td>Economy UV Curing Light</td>
 <td>Henry Schein</td>
 <td>CU-80</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Head-post</td>
 <td>Built in-house</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Mounting bar</td>
 <td>Built in-house</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Mounting block</td>
 <td>Built in-house</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Stereotaxic Frame</td>
 <td>David Kopf Instruments</td>
 <td>902</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Mouse and Neonatal Rat Adaptor</td>
 <td>Stoelting</td>
 <td>51625</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Deltaphase Isothermal Pad</td>
 <td>Braintree Scientific, Inc.</td>
 <td>39DP</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

preparation of an awake mouse for recording neural responses and injecting tracers

It is well known that anesthesia alters neural response properties in various regions of the brain.13. In the auditory system, fundamental response properties of brainstem neurons including threshold, frequency specificity, and inhibitory sidebands are altered in significant ways under anesthesia1-2. These observations prompted physiologists to seek ways to record from single neurons without the contaminating effects of anesthesia. One result was a decerebrate preparation, where the brainstem was completely transected at the level of the midbrain4. The drawbacks of this preparation are a formidable surgery, the elimination of descending projections from the forebrain, and an inability to use sensory stimulation to examine structures above the midbrain. A different strategy has been to implant electrode arrays chronically to record from single neurons and multiunit clusters while the animal is awake and/or behaving5,6. These techniques however are not compatible with injecting tracer dyes after first electrophysiologically characterizing a brain structure. To avoid altering neural response properties with anesthetics while recording electrophysiological response properties from single neurons, we have adapted a head restraint technique long used in bats7-9 to mouse10-12. Using this method, we are able to conduct electrophysiological recordings over several days in the unanesthetized mouse. At the end of the recording sessions, we can then inject a dye to reconstruct electrode positions and recording sites or inject a tracer so that pathways to and from the recording loci can be determined. This method allows for well isolated single neuron recordings over multiple days without the use anesthetics.

Watch this Scientific Journal Video about Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers at JoVE.com

Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

It is well known that anesthesia alters neural response properties in various regions of the brain.13.  In the auditory system, fundamental response ...

preparation-an-awake-mouse-for-recording-neural-responses-injecting

Research

JoVE Journal

Neuroscience

19.3K Views.  Johns Hopkins University. Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

Video: Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

Analyzing Responses of Mouse Olfactory Sensory Neurons Using the Air-phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and other social interactions. The electroolfactogram (EOG) recording is an informative, easy to conduct, and reliable method to assay olfactory function at the level of the olfactory epithelium. Since the 1956 description of the EOG by Ottoson in frogs1, the EOG recording has been applied in many vertebrates including salamanders, rabbits, rats, mice, and humans (reviewed by Scott and Scott-Johnson, 2002, ref. 2). The recent advances in genetic modification in mice have rekindled interest in recording the EOG for physiological characterization of olfactory function in knock-out and knock-in mice. EOG recordings have been successfully applied to demonstrate the central role of olfactory signal transduction components3-8, and more recently to characterize the contribution of certain regulatory mechanisms to OSN responses9-12.
Odorant detection occurs at the surface of the olfactory epithelium on the cilia of OSNs, where a signal transduction cascade leads to opening of ion channels, generating a current that flows into the cilia and depolarizes the membrane13. The EOG is the negative potential recorded extracellularly at the surface of the olfactory epithelium upon odorant stimulation, resulting from a summation of the potential changes caused by individual responsive OSNs in the recording field2. Comparison of the amplitude and kinetics of the EOG thus provide valuable information about how genetic modification and other experimental manipulations influence the molecular signaling underlying the OSN response to odor. 
Here we describe an air-phase EOG recording on a preparation of mouse olfactory turbinates. Briefly, after sacrificing the mouse, the olfactory turbinates are exposed by bisecting the head along the midline and removing the septum. The turbinate preparation is then placed in the recording setup, and a recording electrode is placed at the surface of the olfactory epithelium on one of the medial turbinates. A reference electrode is electrically connected to the tissue through a buffer solution. A continuous stream of humidified air is blown over the surface of the epithelium to keep it moist. The vapor of odorant solutions is puffed into the stream of humidified air to stimulate the epithelium. Responses are recorded and digitized for further analysis.

The electroolfactogram (EOG) recording is an informative, easy-to-conduct, and reliable way of assessing olfactory function at the level of the olfactory epithelium. This protocol describes a recording setup, mouse tissue preparation, data collection, and basic data analysis.

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and ...

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex. Because of the lack of specific pharmacological agents for these channels, we have taken a genetic approach to manipulating channel expression. We use an organotypic culture preparation (16) in order to maintain cell morphology and the laminar pattern of cortex. We typically isolate acute neocortical slices at postnatal days 8-10 and maintain the slices in culture for 3-7 days. This allows us to study neurons at a similar age to those in our work with acute slices and minimizes the development of exuberant excitatory connections in the slice. We record from visually-identified pyramidal neurons in layers II/III or V using infrared illumination (IR-) and differential interference contrast microscopy (DIC) with whole cell patch clamp in current- or voltage-clamp. We use biolistic (Gene gun) transfection of wild type or mutant potassium channel DNA to manipulate expression of the channels to study their function. The transfected cells are easily identified by epifluorescence microscopy after co-transfection with cDNA for green fluorescent protein (GFP). We compare recordings of transfected cells to adjacent, untransfected neurons in the same layer from the same slice.

This is a protocol to prepare and maintain a neocortical slice preparation in organotypic culture for the purpose of making electrical recordings from pyramidal neurons.

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex.  Because of the ...

Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. Here we present a visualized experimental and methodological protocol to directly visualize microregional tissue hypoxia and to infer perivascular oxygen gradients in the mouse cortex. It is based on the non-linear relationship between nicotinamide adenine dinucleotide (NADH) endogenous fluorescence intensity and oxygen partial pressure in the tissue, where observed tissue NADH fluorescence abruptly increases at tissue oxygen levels below 10 mmHg1. We use two-photon excitation at 740 nm which allows for concurrent excitation of intrinsic NADH tissue fluorescence and blood plasma contrasted with Texas-Red dextran. The advantages of this method over existing approaches include the following: it takes advantage of an intrinsic tissue signal and can be performed using standard two-photon in vivo imaging equipment; it permits continuous monitoring in the whole field of view with a depth resolution of ~50 &mu;m. We demonstrate that brain tissue areas furthest from cerebral blood vessels correspond to vulnerable watershed areas which are the first to become functionally hypoxic following a decline in vascular oxygen supply. This method allows one to image microregional cortical oxygenation and is therefore useful for examining the role of inadequate or restricted tissue oxygen supply in neurovascular diseases and stroke.

Here we describe a method to directly visualize microregional tissue hypoxia in the mouse cortex in vivo. It is based on concurrent two-photon imaging of nicotinamide adenine dinucleotide (NADH) and the cortical microcirculation. This method is useful for high resolution analysis of tissue oxygen supply.

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory encoding. LTP elicited in the CA1 region of acute hippocampal slices has been extensively studied. However the molecular mechanisms underlying the maintenance phase of this phenomenon are still poorly understood. This could be partly due to the various experimental conditions used by different laboratories. Indeed, the maintenance phase of LTP is strongly dependent on external parameters like oxygenation, temperature and humidity. It is also dependent on internal parameters like orientation of the slicing plane and slice viability after dissection.
The optimization of all these parameters enables the induction of a very reproducible and very stable long-term potentiation. This methodology offers the possibility to further explore the molecular mechanisms involved in the stable increase in synaptic strength in hippocampal slices. It also highlights the importance of experimental conditions in in vitro investigation of neurophysiological phenomena.

This paper presents a complete methodology to prepare and preserve in vitro acute hippocampal slices from adult mice. This protocol allows the recording of very stable long-lasting long-term potentiation (LTP) for more than 8 hours with a success rate of 95%.

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory ...

Neural Circuit Recording from an Intact Cockroach Nervous System

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological effects of insecticides. This article describes the scope of cockroach sensory modalities that can be used to assay how an insect nervous system responds to environmental perturbations. Emphasis here is on the escape behavior mediated by cerci to giant fiber transmission in Periplaneta americana. This in situ preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of neuronal activity. Peptides or other chemical reagents can then be applied directly to the nervous system in solution with the physiological saline. Insecticides could also be administered prior to dissection and the escape circuit can serve as a proxy for the excitable state of the central nervous system. In this context the assays described herein would also be useful to researchers interested in limb regeneration and the evolution of nervous system development for which P. americana is an established model organism.

This article describes the cockroach ventral nerve cord dissection and extracellular recordings from the cercal nerve and connectives. Evoked responses are generated by electrical stimulation of the cercal nerve or direct mechanical stimulation of the cerci.

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

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 ...

Fabrication of High Contact-Density, Flat-Interface Nerve Electrodes for Recording and Stimulation Applications

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic applications. This protocol describes a fabrication technique of a modified cylindrical nerve cuff electrode to meet these criteria. Minimum computer-aided design and manufacturing (CAD and CAM) skills are necessary to consistently produce cuffs with high precision (contact placement 0.51 &#177; 0.04 mm) and various cuff sizes. The precision in spatially distributing the contacts and the ability to retain a predefined geometry accomplished with this design are two criteria essential to optimize the cuff&#39;s interface for selective recording and stimulation. The presented design also maximizes the flexibility in the longitudinal direction while maintaining sufficient rigidity in the transverse direction to reshape the nerve by using materials with different elasticities. The expansion of the cuff&#39;s cross sectional area as a result of increasing the pressure inside the cuff was observed to be 25% at 67 mm Hg. This test demonstrates the flexibility of the cuff and its response to nerve swelling post-implant. The stability of the contacts&#39; interface and recording quality were also examined with contacts&#39; impedance and signal-to-noise ratio metrics from a chronically implanted cuff (7.5 months), and observed to be 2.55 &#177; 0.25 k&#8486; and 5.10 &#177; 0.81 dB respectively.

This article provides a detailed description on the fabrication process of a high contact-density flat interface nerve electrode (FINE). This electrode is optimized for recording and stimulating neural activity selectively within peripheral nerves.

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic ...

Construction of an Improved Multi-Tetrode Hyperdrive for Large-Scale Neural Recording in Behaving Rats

Monitoring the activity patterns of a large population of neurons over many days in awake animals is a valuable technique in the field of systems neuroscience. One key component of this technique consists of the precise placement of multiple electrodes into desired brain regions and the maintenance of their stability. Here, we describe a protocol for the construction of a 3D-printable hyperdrive, which includes eighteen independently adjustable tetrodes, and is specifically designed for in vivo extracellular neural recording in freely behaving rats. The tetrodes attached to the microdrives can either be individually advanced into multiple brain regions along the track, or can be used to place an array of electrodes into a smaller area. The multiple tetrodes allow for simultaneous examination of action potentials from dozens of individual neurons, as well as local field potentials from populations of neurons in the brain during active behavior. In addition, the design provides for simpler 3D drafting software that can easily be modified for differing experimental needs.

We present the construction of a 3D-printable hyperdrive with eighteen independently adjustable tetrodes. The hyperdrive is designed to record brain activity in freely behaving rats over a period of several weeks.