We are very grateful to Wenbiao Gan (Skirball Institute, Department of Physiology and Neuroscience, New York University School of Medicine, USA) for excellent technical guidance and Patti Raley, Medical Editor (Department of Neurological Surgery, Indiana University School of Medicine), for editing the manuscript. We thank Dr. Jinhui Chen (Stark Neurosciences Research Institute, Indiana University School of Medicine, USA) for providing B6.Cg-Tg(Thy1-YFP)HJrs/J mice. This work was supported in part by the Foundation of Director of General Hospital of Jinan Military Region of Chines PLA 2016ZD03 (XJL), and NIH NS059622, NS073636, DOD CDMRP W81XWH-12-1-0562, Merit Review Award I01 BX002356 from the U.S. Department of Veterans Affairs, Craig H Neilsen Foundation 296749, Indiana Spinal Cord and Brain Injury Research Foundation, Mari Hulman George Endowment Funds (XMX).

All surgical and animal handling procedures were performed as approved under the Guide for the Care and Use of Laboratory Animals (National Research Council) and the Guidelines of the Indiana University School of Medicine Institutional Animal Care and Use Committee. The schematic illustration of the 2-photon imaging setup is shown in Figure 1.
1. Preparing the Animal for Imaging
<ol>
	<li>Place sterile gauze, cotton swabs, and autoclaved surgical tools in the surgery area (Table 1). Turn on a Micro Bead Sterilizer for intersurgery sterilization of surgical tools.</li>
	<li>Anesthetize the mouse (male or female, 1.5&#8211;2.5 months, 20&#8211;25 g) by intraperitoneal injection (i.p.) of a mixture of ketamine (17.2 mg/mL), xylazine (0.475 mg/mL) and acepromazine (0.238 mg/mL) (6 &#181;L/g body weight). Wait until the proper depth of anesthesia is reached when the animal ceases to respond to a toe pinch stimulus and has no corneal reflex. During the surgery, give a booster dose of 1/3 the original dose of the anesthetic cocktail as needed to maintain the original anesthetic state.</li>
	<li>Inject buprenorphine subcutaneously (s.c.; 0.05&#8211;0.10 mg/kg) as an analgesic agent prior to surgery. Apply protective ointment to the eyes of the animal to prevent corneal drying and cataract formation.</li>
	<li>Place a GCaMP6s transgenic mouse on a heating pad to prevent hypothermia and cover it with a sterile surgical drape. Stabilize the animal&#39;s head with a Head Holding Adaptor for Mice.</li>
	<li>Shave the head over most of the scalp with a double-edge razor blade. Clean the surgical area with a sterile alcohol preparation pad followed by povidone-iodine solution.</li>
</ol>
2. Chronic Cranial Window Preparation
<ol>
	<li>Make a rectangular cut of the scalp (2 mm x 3 mm) with a pair of scissors on one side (right) of the skull. Push the skin aside with a cotton swab to create an exposure area of &#62;3 mm in diameter (Figure 3A). Remove the connective tissue attached to the skull by gently scraping the skull with a blunt microsurgical blade (Scalpel Blade #10; Figure 3C).</li>
	<li>Mark the S1 area by gently drawing a circle (3 mm in diameter, center coordination: -1.5 mm from the bregma and 2.5 mm from the midline) on the skull using a dental drill (Figure 2B).</li>
	<li>Perform the same procedure on the left skull (Figure 2B).</li>
	<li>Apply a thin layer of Cyanoacrylate Super Glue to the bone to provide a base for dental cement application.</li>
	<li>Under a dissecting microscope, thin down a circular groove in the skull around the S1 area using a high-speed microdrill (Figure 3D). The purpose is to create a smooth edge for placement of an optical window onto it.</li>
	<li>Repeatedly apply artificial cerebrospinal fluid (ACSF, room temperature) to the skull periodically during the thinning process to facilitate drilling and dissipate heat. Perform the drilling intermittently to avoid friction-induced overheating. Suck away the bone debris with a house vacuum line or a vacuum pump.</li>
	<li>After approximately 2/3 depth of the bone is drilled, slowly and carefully thin the remaining 1/3 bone until a circular piece of the skull (bone flap) is completely free from the surrounding skull.</li>
	<li>Slowly and carefully remove the circular bone flap with a pair of # 5/45 forceps and expose the dura (Figure 2C; Figure 3E).</li>
	<li>Keep the exposed brain region moist with ACSF (Figure 3E). Avoid any damage to pial vessels, since hemorrhage will alter cerebral blood flow, accelerate brain swelling, and severely degrade imaging quality.</li>
	<li>Perform the same procedure on the left (contralateral) side of the skull.</li>
	<li>For assembling the optical window, use optical adhesive to glue and cure between coverglass layers one at a time. If necessary, warm the optical window (50 &#176;C for 12 h) in an incubator to enhance the adhesive bonding. The optical window contains 2 portions: the top portion contains a single round cover glass with a diameter of 5 mm and the bottom portion contains 1&#8211;3 round cover glasses with a diameter of 3 mm (Figure 2D).</li>
	<li>Store the optical window in ethanol (70%, vol/vol) and rinse it with sterile saline before use. Before optical window installation, check the optical window for imperfections, including an incorrect amount of optical adhesive or inaccurate alignment under a stereoscope.</li>
	<li>Install the optical window over the craniotomy region (Figure 2D). The top portion of the optical window rests on the skull and the bottom portion of the optical window fits within the craniotomized opening and rests on the dura in the presence of CSF (Figure 2D, E).</li>
	<li>Seal the top portion of the optical window edge to the skull with the Cyanoacrylate Super Glue (Figure 2F; Figure 3F).</li>
	<li>When the Cyanoacrylate Super Glue has dried, apply black dental cement (Dentsply) to cover the glass edge. Apply dental cement to all the exposed skull and wound margins to block light (Figure 2G-H; Figure 3B).</li>
	<li>Perform the same procedure on the left side.</li>
	<li>Allow the animal to recover on the heating pad and return the animal to its home cage for recovery under appropriate monitoring. Provide wet food to facilitate chewing and hydration. To reduce pain, administer buprenorphine s.c. (0.05&#8211;2.0 mg/kg) every 8&#8211;12 h postinjury for 2 days. Allow the mouse to recover for an additional 7&#8211;10 days before imaging.</li>
</ol>
3. Two-photon imaging
<ol>
	<li>On the day of the experiment, anesthetize the animal with ketamine and xylazine (doses and maintenance of anesthesia as described above). Clean the cranial window with 75% (vol/vol) ethanol.</li>
	<li>Place the mouse under an imaging set-up consisting of a 2P microscope and the Head Holding Adaptor, resting on a heating pad with its head immobilized through ear bars attached to a mouse head holder.</li>
	<li>Image one side of the optical window at a time. To minimize optical aberrations, make sure that the optical window and right skull are oriented perpendicular to the optical axis of the microscope.</li>
	<li>Take an initial picture of the cranial window with bright field illumination at 4x magnification as a reference map for registration with higher magnification 2P angiographies (Figure 4A1).</li>
	<li>Zoom in on the area of interest using 10x magnification (Figure 4A2).</li>
	<li>Select an area of interest in the S1 area in cortical layers I or II (up to 150 &#181;m below the pial surface). If higher magnification is desired, use a 20X objective.</li>
	<li>Search for a field of view with multiple neurons. Acquire images using a 2P microscope (Figure 4B). Determine the exposure time and excitation light level (&#8776;910 nm) based on the depth and expression level of GCaMP6s, with weaker signals requiring longer exposure times and concordantly less time resolution. We typically use ~4 &#181;s per pixel exposure times. Acquire images at frame rates of 3&#8211;5 Hz at a resolution of 512 &#215; 512 pixels.</li>
	<li>Start to record the time series. Store coordinates of every region of interest (ROI) relative to the vascular pattern or to fiducial points in low magnification images. Image the same ROI over time. Perform the same procedure to observe calcium dynamics of the S1 area on the left hemisphere. Calculate changes in fluorescence from each ROI.</li>
	<li>Acquire images either individually or as a time-lapse movie (Figure 4C1-C3, D, E).</li>
	<li>Study cortical blood flow dynamics in vivo by imaging fluorescent dextran dyes injected into the tail vein of rodents with 2P microscopy. Use vasculatures as landmarks to recognize and image the same ROI in imaging sessions over long periods.</li>
</ol>
4. Data Processing
<ol>
	<li>Use ImageJ and Origin for image analysis. Import image sequences with ImageJ (Figure 5A-B). For individual images (or t-series movies), select an ROI within an imaged neuron and 2 separate nearby regions for background measurement (Figure 5C). Acquire total intensity with an ROI manager (Figure 5C). 
	NOTE: Figure 6 shows representative calcium images. Calcium transient (green) and blood vessel (red) show at different time points in seconds (Figure 6A-D) in a mouse at 7 d after the initial window installation. The z-projection of a 3 min recording period for red and green channels (Figure 6E) or green channel (Figure 6F) is also shown. A z-projection displays a compressed image of all images in the movie from which a rough and circle boundary (encompassing the object neuron of interest in all frames) is manually selected as well as the selection of the 2 independent nearby background regions (Figure 6G).</li>
	<li>From each ROI, calculate the average fluorescence background FB, FB = (FB1+FB2)/2, and the fluorescence changes of the soma (F) expressed as &#916;F/F, &#916;F/F = (F-FB)/FB.</li>
	<li>Perform the peak analysis step by step using functions of ImageJ including Baseline Correction, Peak Finding/Determination, Peak Integration, Peak Fitting, and/or Time-Saving Peak Analysis Features.</li>
</ol>

<ol>
	<li>Cichon, J., Gan, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Branch-specific+dendritic+Ca(2+)+spikes+cause+persistent+synaptic+plasticity.">Branch-specific dendritic Ca(2+) spikes cause persistent synaptic plasticity.</a> Nature. 520 (7546), 180-185 (2015).</li><li>Andermann, M. L., Kerlin, A. M., Roumis, D. K., Glickfeld, L. L., Reid, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+specialization+of+mouse+higher+visual+cortical+areas.">Functional specialization of mouse higher visual cortical areas.</a> Neuron. 72 (6), 1025-1039 (2011).</li><li>Chen, X., Leischner, U., Rochefort, N. L., Nelken, I., Konnerth, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+mapping+of+single+spines+in+cortical+neurons+in+vivo.">Functional mapping of single spines in cortical neurons in vivo.</a> Nature. 475 (7357), 501-505 (2011).</li><li>Mittmann, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+calcium+imaging+of+evoked+activity+from+L5+somatosensory+neurons+in+vivo.">Two-photon calcium imaging of evoked activity from L5 somatosensory neurons in vivo.</a> Nat Neurosci. 14 (8), 1089-1093 (2011).</li><li>Katona, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+two-photon+in+vivo+imaging+with+three-dimensional+random-access+scanning+in+large+tissue+volumes.">Fast two-photon in vivo imaging with three-dimensional random-access scanning in large tissue volumes.</a> Nat Methods. 9 (2), 201-208 (2012).</li><li>Zhang, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Remodeling+the+Dendritic+Spines+in+the+Hindlimb+Representation+of+the+Sensory+Cortex+after+Spinal+Cord+Hemisection+in+Mice.">Remodeling the Dendritic Spines in the Hindlimb Representation of the Sensory Cortex after Spinal Cord Hemisection in Mice.</a> PLoS One. 10 (7), e0132077 (2015).</li><li>Keck, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+scaling+and+homeostatic+plasticity+in+the+mouse+visual+cortex+in+vivo.">Synaptic scaling and homeostatic plasticity in the mouse visual cortex in vivo.</a> Neuron. 80 (2), 327-334 (2013).</li><li>Helmchen, F., Fee, M. 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Authors have nothing to disclose.

Two-photon imaging enables the simultaneous measurement of the activity of many cells at a reasonable resolution at up to approximately 800 &#181;m below the brain surface. We integrated 2P and a dual optical window technique with genetically encoded GCaMP6s to observe calcium dynamics in populations of layer V neuronal somata located &#62;500 &#181;m below the pia. With an open optical window on each side of the skull, the method enables recordings of neural activity across symmetric mammalian brain regions for prolonged and stable calcium imaging in vivo. Key aspects of this method include the performance of minimally invasive surgery to create dual open skull optical windows and 2P imaging of neuronal activity in symmetrical S1 regions.
Examining the network activity in bilateral S1 regions is an example of how dual cranial windows could be used to probe local and symmetric information processing in the adult brain in vivo. This method also can be used to study the neuronal activity (e.g. using Thy1-GCaMP6s mice) and cortical plasticity B6.Cg-Tg(Thy1-YFP)HJrs/J from intact tissue of the corresponding symmetric regions in recovery after deafferentation due to a unilateral CNS injury. The method can also be used to examine cortical activity in response to peripheral stimulation strategies such as optogenetic, electrical or chemical stimulations. Although we only demonstrate the technique in the S1 area, this approach could be employed to investigate activities in other symmetric areas in the living brain such as the layer V neurons of the primary motor cortex (M1). Observations on reorganization of brain regions may provide a neural substrate for adaptive motor behavior, which plays a critical role in postinjury recovery of function. It also allows for chronic measurement of the symmetrical activity of genetically identified cells during behavior in head-fixed mice.
Technically, investigators should pay careful attention to the quality and consistency of dual cranial windows. Thinned-skull cranial window technique 12 is highly recommended for the beginners to practice. Spines are calcium compartments because of their morphology and local influx and extrusion mechanisms. In this study, we used B6.Cg-Tg(Thy1-YFP)HJrs/J mice as an example to demonstrate dendritic spine dynamics in vivo (Figure 4). An open-skull glass window installation may cause high spine turnover and reactive glial responses after surgery 12. In contrast, with the thinned-skull window technique, spines and dendrites can be well preserved.
Choices of dimensions and thicknesses of optical windows will be dependent on the desired field of view, the skull curvature, skull thickness, and the type of imaging 13. Insufficient pressure of the optical coverglasses on the dura may cause thickening of the dura, regrowth of the skull, and increased brain motion. In contrast, excessive pressure of the optical coverglasses may impede cortical blood flow.
For optical window assembly, glue and cure additional layers of coverglasses one at a time with optical adhesive and long-wavelength UV light. To help strengthen the adhesive bonding, warm the fabricated window (50 &#176;C for 12 h) in an incubator. Store the optical window coverglass in 70% (vol/vol) ethanol and, before the surgery, rinse it with sterile saline. The optical window needs to be examined for imperfections (such as an incorrect amount of optical adhesive or inaccurate alignment) under a stereoscope before use to avoid failure. If the optical adhesive is too thin, air spaces may be creased, which can cause optical aberrations or cover glass detachment at implantation. In contrast, too much optical adhesive can cause overflow past the edges of the cranial window.
In summary, here we describe an experimental procedure for measuring calcium dynamics or dendritic morphology through cranial windows from 2 symmetric primary somatosensory cortical regions in Thy1-GCaMP6s and Thy1-YFP transgenic mice, respectively, using 2P microscopy. This technique may greatly facilitate dissecting the complex structural and functional relationship of neural networks.

Calcium and its regulation are essential in mediating multiple physiological processes. Monitoring calcium dynamics is useful for understanding normal brain function as well as pathological conditions of brain disorders 1,2,3,4,5,6,7,8. Imaging GCaMP6s fluorescence is a powerful tool for quantifying spike numbers, timing, and frequency, as well as levels of synaptic input both in vitro and in vivo 9,10,11.
The mammalian brain exhibits marked symmetry across the sagittal plane. Although recent neurological research has shed light on the cortical remodeling and neuronal activity triggered by traumatic brain or spinal cord injury 6,7, the roles that intact tissue of symmetrically corresponding regions play in recovery are still obscure.
In this study, we describe experimental procedures for creating bilateral symmetric optical windows for in vivo imaging. Using these optical windows, we describe the measuring of calcium dynamics from primary somatosensory cortices (S1) in GCaMP6s transgenic mice using 2P microscopy. In the results section, we also show in vivo dendritic morphology at different time points in the S1 region in the EGFP transgenic mice using 2P imaging. The observation method regarding the network dynamics in the bilateral S1 areas is but an initial example of how dual open cranial windows will help neuroscientists to probe local and symmetric information processing in the adult mammalian brain in normal conditions or in different disease models in vivo.

<table border="0" cellpadding="0" cellspacing="0" width="642">
	<tbody>
		<tr height="81">
			<td height="81" style="height:81px;width:208px;">C57BL/6J-Tg(Thy1-GCaMP6s)GP4.3Dkim/J</td>
			<td style="width:103px;">Jackson</td>
			<td align="right" style="width:83px;">24275</td>
			<td style="width:249px;">Can be any strains made by the genetically-encoded neuronal indicator and effector expressing the green fluorescent calcium indicator, GCaMP, in subsets of excitatory neurons in the cortex.</td>
		</tr>
		<tr height="65">
			<td height="65" style="height:65px;width:208px;">B6.Cg-Tg(Thy1-EGFP)OJrs/GfngJ.</td>
			<td style="width:103px;">Jackson</td>
			<td align="right" style="width:83px;">7919</td>
			<td style="width:249px;">Can be any strains expressing EGFP in subsets of neurons within specific populations in the cortex; providing a bright, vital Golgi-like stain.&#160;</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Dumont no. 5 forceps, standard, Dumoxel</td>
			<td style="width:103px;">Fine Science Tools</td>
			<td style="width:83px;">11251-30</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Dumont no. 5SF forceps</td>
			<td style="width:103px;">Fine Science Tools</td>
			<td style="width:83px;">11252-00</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Micro Bead Sterilizer</td>
			<td style="width:103px;">Southern Labware</td>
			<td style="width:83px;">B1201</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Harishige&#8217;s SG-4N Head Holding Adaptor</td>
			<td style="width:103px;">Tritech Research</td>
			<td style="width:83px;">SG-4N</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Ideal Micro-Drill Complete Kit</td>
			<td style="width:103px;">Harvard Apparatus</td>
			<td style="width:83px;">72-6065</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Burrs for Micro Drill</td>
			<td style="width:103px;">Fine Science Tools</td>
			<td style="width:83px;">19007-05</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Round cover glass, 3 mm</td>
			<td style="width:103px;">Harvard Biosciences</td>
			<td style="width:83px;">W4 64-0720</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;width:208px;">Round cover glass, 5 mm</td>
			<td style="width:103px;">Harvard Biosciences</td>
			<td style="width:83px;">W4 64-0700</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:208px;">Norland optical adhesive</td>
			<td style="width:103px;">Norland Products</td>
			<td align="right" style="width:83px;">7106</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:208px;">Loctite liquid super glue</td>
			<td style="width:103px;">WB Mason</td>
			<td style="width:83px;">LOC1647358</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="81">
			<td height="81" style="height:81px;width:208px;">Grip cement kit, powder and solvent</td>
			<td style="width:103px;">Dentsply</td>
			<td align="right" style="width:83px;">675570</td>
			<td style="width:249px;">Can be any brand of choice. Mix 5 parts of white dental cement powder (Grip Cement) with one part of black Tempera powder paint (to shield from light; some users prefer a 10:1 ratio).</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:208px;">Gel foam</td>
			<td style="width:103px;">Moore Medical</td>
			<td align="right" style="width:83px;">2928</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:208px;">Micro dissecting scissors</td>
			<td style="width:103px;">Roboz</td>
			<td style="width:83px;">RS-5841</td>
			<td style="width:249px;">Can be any brand of choice.</td>
		</tr>
		<tr height="113">
			<td height="113" style="height:113px;width:208px;">Artificial cerebrospinal fluid (ASF)</td>
			<td style="width:103px;">-</td>
			<td style="width:83px;">-</td>
			<td style="width:249px;">The ACSF contains 125 mM NaCl, 4.5 mM KCl, 26 mM NaHCO3, 1.25 mM NaH2PO4, 2 mM CaCl2, 1 mM MgCl2 and 20 mM glucose. Bubble the ACSF with oxygen (95% oxygen, 5% carbon dioxide) and adjust the pH to 7.4. Prepare fresh ACSF solution before every experiment.</td>
		</tr>
	</tbody>
</table>

imaging neural activity in the primary somatosensory cortex using thy1-gcamp6s transgenic mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Using the bilateral optical window, we obtained results from 2 separate experiments. The first experiment employed EGFP-expressing mice to image dendritic varicosities. Figure 4C1-3 shows examples of images captured at different time points after optical window implantation. Note that the number and location of the dendritic branches and spines are remarkably stable between the 2 views (Figure 4D, E, z-projection).
The second experiment employed Thy1-GCaMP6s transgenic mice to measure the intracellular calcium transient, thus illustrating how calcium physiology can be measured within a single neuron in vivo (Figure 6). This technique can be used to record and quantify spontaneous activity in populations of layer V pyramidal neurons located at as deep as &#62;500 &#181;m below the pia. Average spontaneous responses over time (calculated as the average responses at each time point) can be calculated across numerous trials. The 2P technique has the advantage of detecting activity simultaneously in large or dispersed neuronal populations over an extended period with little mechanical disturbance to the brain as compared to multielectrode recordings.
To generate an acceptable average measurement, 5&#8211;10 trials are recommended. The number of trials will be dependent on the spontaneous responses or inherent variability of the responses to a particular stimulation. If all the steps are followed strictly, as described above, a researcher trained in both thinned-skull cranial window technique and in vivo 2P calcium imaging should be able to obtain recordings of 100&#8211;200 neurons on both sides from 1&#8211;2 animals per day.
After the peak analysis, a collection of results such as the real location of the peak, the real amplitude, the area, and the full width at half maximum (FWHM) for each peak can be obtained.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56297/56297fig1.jpg" /> 
Figure 1: Schematic illustration of the components of a cranial optical window for 2-photon (2P) imaging. <a href="https://cloudflare.jove.com/files/ftp_upload/56297/56297fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56297/56297fig2.jpg" /> 
Figure 2: Schematic drawing shows key steps for preparing a cranial window.&#160;(A) Under a dissecting microscope, remove the skin over the surgical region using a pair of surgical scissors. (B) Use a high-speed microdrill to produce a circular groove (3 mm diameter) on the skull over the S1 area until the bone flap within the groove becomes detached from the surrounding bone. (C) Slowly remove the bone flap to expose the underlying dura. (D) Assemble the optical window with coverglasses with 2 different diameters. The upper coverglass has a larger diameter (5 mm) and the lower coverglasses (usually 1-3) have a smaller diameter (3 mm). The thickness of the coverglasses to be installed will be dependent on the thickness of the skull. (E) Place the coverglass onto the circular opening of the skull to create an optical window. The lower portion of the coverglasses should fit within the cranial opening. The bottom of the coverglasses should gently contact the dura. (F) Seal the window edges to the skull with cyanoacrylate glue. (G) When the cyanoacrylate dries, apply black dental cement lateral to the glue wall. (H) Fill the cranial window with ddH2O and use water-immersion objectives for imaging. <a href="https://cloudflare.jove.com/files/ftp_upload/56297/56297fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56297/56297fig3.jpg" /> 
Figure 3. Creation of bilateral windows on the skull.&#160;(A) A photographic image shows 2 cranial optical windows on each side of the skull overlying the S1 area. (B) High magnification of the boxed region of image (A) showing bilateral optical windows exposing the underlying dura and blood vessels. Scale bar = 680 &#181;m. (C) The exposed skull. (D) Creation of a circular groove within which a central bone flap is seen. Scale bar = 900 &#181;m. (E) After the removal of the bone flap, the cranial opening was filled with artificial cerebrospinal fluid (ACSF). Scale bar = 720 &#181;m. (F) Installation of the optical window over the cranial opening and sealing the edge of the window with super glue. The dura and blood vessels are visible through the optical window. Scale bar = 600 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56297/56297fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56297/56297fig4.jpg" /> 
Figure 4: Calcium and dendritic imaging.&#160;(A1) Identify regions of interest using blood vessels (BV) as landmarks in Thy1-GCaMP6s mice. (A2) Identify the same labeled cells (red arrows) over time using the same landmarks and record. (B) Representative Z-projection of calcium signals in neurons (red arrows) of the primary somatosensory cortex (S1). Scale bar = 10 &#181;m. (C1-3) Visualization of dendrites (green) and a blood vessel (red) in B6.Cg-Tg(Thy1-YFP)HJrs/J at different depths in the layer II of the S1 region at 1 day after optical window installation. (D) Z-projection of multiple images at the same region at 1 day. (E) Z-projection of multiple images at the same region at 7 days after the optical window installation. Note the remarkable stability of the number and location of adult spines and dendritic branches between 1 day and 7 days after optical window installation. Scale bar = C-E 5 &#181;m (C-E). <a href="https://cloudflare.jove.com/files/ftp_upload/56297/56297fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a><img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56297/56297fig5.jpg" /> 
Figure 5: Calcium signal tracing.&#160;(A, B) Examples of imported t-series movies with ImageJ for data analysis of 2 neurons (neuron 1 and neuron 2, respectively). (C) A spreadsheet shows data on illumination intensity from region of interest (ROI; within the neuron and 2 separate nearby regions as background measures). (D) Calculation of the peak: &#916;F/F= (F-FB)/FB, FB = (FB1+FB2)/2 (the average fluorescence background [FB], and the fluorescence changes of the soma [F] expressed as &#916;F/F). <a href="https://cloudflare.jove.com/files/ftp_upload/56297/56297fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56297/56297fig6.jpg" /> 
Figure 6. Representative calcium images.&#160;(A-D) Calcium transient (green) and blood vessels (red) at different time points in seconds in a mouse at 7 days after the optical window installation. Yellow arrow: a relatively less active neuron. White arrow: a spiking neuron. (E, F) Z-projection of a 3 min recording period for the red (blood vessels) and green (calcium) channels (E) and green channel only (F). (G) Selected neurons and their adjacent background areas are marked. Scale bar = 10 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56297/56297fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice at JoVE.com

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

imaging-neural-activity-primary-somatosensory-cortex-using-thy1

Nanyang Technological University

Research

JoVE Journal

Neuroscience

10.9K Views.  Indiana University School of Medicine. We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

Video: Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

High-density EEG Recordings of the Freely Moving Mice using Polyimide-based Microelectrode

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a noninvasive brain monitoring tool in cognitive neuroscience as well as a diagnostic tool for epilepsy and sleep disorders in neurology. However, the underlying mechanism of EEG rhythm generation is still under the veil. Recently introduced polyimide-based microelectrode (PBM-array) for high resolution mouse EEG1 is one of the trials to answer the neurophysiological questions on EEG signals based on a rich genetic resource that the mouse model contains for the analysis of complex EEG generation process. This application of nanofabricated PBM-array to mouse skull is an efficient tool for collecting large-scale brain activity of transgenic mice and accommodates to identify the neural correlates to certain EEG rhythms in conjunction with behavior. However its ultra-thin thickness and bifurcated structure cause a trouble in handling and implantation of PBM-array. In the presented video, the preparation and surgery steps for the implantation of PBM-array on a mouse skull are described step by step. Handling and surgery tips to help researchers succeed in implantation are also provided.

In this article, we described the surgery procedure and handling tips for implantation of ultra-thin polyimide-based microelectrode array (PBM-array) on the mouse skull for acquisition of high-density encephalography (EEG) in a mouse model.

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a ...

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. ...

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 ...

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

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 ...

Bioluminescence Imaging of Neuroinflammation in Transgenic Mice After Peripheral Inoculation of Alpha-Synuclein Fibrils

To study the prion-like behavior of misfolded alpha-synuclein, mouse models are needed that allow fast and simple transmission of alpha-synuclein prionoids, which cause neuropathology within the central nervous system (CNS). Here we describe that intraglossal or intraperitoneal injection of alpha-synuclein fibrils into bigenic Tg(M83+/-:Gfap-luc+/-) mice, which overexpress human alpha-synuclein with the A53T mutation from the prion protein promoter and firefly luciferase from the promoter for glial fibrillary acidic protein (Gfap), is sufficient to induce neuropathologic disease. In comparison to homozygous Tg(M83+/+) mice that develop severe neurologic symptoms beginning at an age of 8 months, heterozygous Tg(M83+/-:Gfap-luc+/-) animals remain free of spontaneous disease until they reach an age of 22 months. Interestingly, injection of alpha-synuclein fibrils via the intraperitoneal route induced neurologic disease with paralysis in four of five Tg(M83+/-:Gfap-luc+/-) mice with a median incubation time of 229 &#177;17 days. Diseased animals showed severe deposits of phosphorylated alpha-synuclein in their brains and spinal cords. Accumulations of alpha-synuclein were sarkosyl-insoluble and colocalized with ubiquitin and p62, and were accompanied by an inflammatory response resulting in astrocytic gliosis and microgliosis. Surprisingly, inoculation of alpha-synuclein fibrils into the tongue was less effective in causing disease with only one of five injected animals showing alpha-synuclein pathology after 285 days. Our findings show that inoculation via the intraglossal route and more so via the intraperitoneal route is suitable to induce neurologic illness with relevant hallmarks of synucleinopathies in Tg(M83+/-:Gfap-luc+/-) mice. This provides a new model for studying prion-like pathogenesis induced by alpha-synuclein prionoids in greater detail.

Peripheral injection of alpha-synuclein fibrils into the peritoneum or tongue of Tg(M83+/-:Gfap-luc+/-) mice, which express human alpha-synuclein with the familial A53T mutation and firefly luciferase, can induce neuropathology, including neuroinflammation, in their central nervous system.

To study the prion-like behavior of misfolded alpha-synuclein, mouse models are needed that allow fast and simple transmission of alpha-synuclein ...

Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or postmitotic neurons. Later, some progenitors switch to a gliogenic fate, adding to the astrocyte and oligodendrocyte populations. Using time-lapse video-microscopy of primary cerebral cortex cell cultures, it is possible to study the cellular and molecular mechanisms controlling the mode of cell division and cell cycle parameters of progenitor cells. Similarly, the fate of postmitotic cells can be examined using cell-specific fluorescent reporter proteins or post-imaging immunocytochemistry. More importantly, all these features can be analyzed at the single-cell level, allowing the identification of progenitors committed to the generation of specific cell types. Manipulation of gene expression can also be performed using viral-mediated transfection, allowing the study of cell-autonomous and non-cell-autonomous phenomena. Finally, the use of fusion fluorescent proteins allows the study of symmetric and asymmetric distribution of selected proteins during division and the correlation with daughter cells fate. Here, we describe the time-lapse video-microscopy method to image primary cerebral cortex murine cells for up to several days and analyze the mode of cell division, cell cycle length and fate of newly generated cells. We also describe a simple method to transfect progenitor cells, which can be applied to manipulate genes of interest or simply label cells with reporter proteins.

Live imaging is a powerful tool to study cellular behaviors in real time. Here, we describe a protocol for time-lapse video-microscopy of primary cerebral cortex cells that allows a detailed examination of the phases enacted during the lineage progression from primary neural stem cells to differentiated neurons and glia.

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or ...

Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral tissues. The hypothalamus in the central nervous system (CNS) is the control center for energy and insulin signal response regulation. Chronic inflammation in peripheral tissues and imbalances of certain chemokines (such as CCL5, TNF&#945;, and IL-6) contribute to diabetes and obesity. However, the functional mechanism(s) connecting chemokines and hypothalamic insulin signal regulation still remain unclear.
In vitro primary neuron culture models are convenient and simple models which can be used to investigate insulin signal regulation in hypothalamic neurons. In this study, we introduced exogeneous GLUT4 protein conjugated with GFP (GFP-GLUT4) into primary hypothalamic neurons to track GLUT4 membrane translocation upon insulin stimulation. Time-lapse images of GFP-GLUT4 protein trafficking were recorded by deconvolution microscopy, which allowed users to generate high-speed, high-resolution images without damaging the neurons significantly while conducting the experiment. The contribution of CCR5 in insulin regulated GLUT4 translocation was observed in CCR5 deficient hypothalamic neurons, which were isolated and cultured from CCR5 knockout mice. Our results demonstrated that the GLUT4 membrane translocation efficiency was reduced in CCR5 deficient hypothalamic neurons after insulin stimulation.

This protocol describes a technique for observation of real-time Green Fluorescence Protein (GFP) tagged Glucose Transporter 4 (GLUT4) protein trafficking upon insulin stimulation and characterization of the biological role of CCR5 in the insulin&#8211;GLUT4 signaling pathway with Deconvolution Microscopy.

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral ...

Analyzing Neural Activity and Connectivity Using Intracranial EEG Data with SPM Software

Measuring neural activity and connectivity associated with cognitive functions at high spatial and temporal resolutions is an important goal in cognitive neuroscience. Intracranial electroencephalography (EEG) can directly record electrical neural activity and has the unique potential to accomplish this goal. Traditionally, averaging analysis has been applied to analyze intracranial EEG data; however, several new techniques are available for depicting neural activity and intra- and inter-regional connectivity. Here, we introduce two analytical protocols we recently applied to analyze intracranial EEG data using the Statistical Parametric Mapping (SPM) software: time-frequency SPM analysis for neural activity and dynamic causal modeling of induced responses for intra- and inter-regional connectivity. We report our analysis of intracranial EEG data during the observation of faces as representative results. The results revealed that the inferior occipital gyrus (IOG) showed gamma-band activity at very early stages (110 ms) in response to faces, and both the IOG and amygdala showed rapid intra- and inter-regional connectivity using various types of oscillations. These analytical protocols have the potential to identify the neural mechanisms underlying cognitive functions with high spatial and temporal profiles.

We present two analytical protocols that can be used to analyze intracranial electroencephalography data using the Statistical Parametric Mapping (SPM) software: time-frequency statistical parametric mapping analysis for neural activity, and dynamic causal modeling of induced responses for intra- and inter-regional connectivity.

Measuring neural activity and connectivity associated with cognitive functions at high spatial and temporal resolutions is an important goal in cognitive ...

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. Adeno-associated viruses (AAVs) are increasingly used to improve understanding of brain circuits and their functions. The AAV-mediated manipulation of specific cell populations and/or of precise molecular components has been tremendously useful to identify new sleep regulatory circuits/molecules and key proteins contributing to the adverse effects of sleep loss. For instance, inhibiting activity of the filamentous actin-severing protein&#160;cofilin&#160;using AAV prevents sleep deprivation-induced memory impairment. Here, a protocol is&#160;described that combines the manipulation of cofilin function in a cerebral cortex area with the recording of ECoG activity to examine whether cortical cofilin modulates the wakefulness and sleep ECoG signals. AAV injection is performed during the same surgical procedure as the implantation of ECoG and electromyographic (EMG) electrodes in adult male and female mice. Mice are anesthetized, and their heads are shaved. After skin cleaning and incision, stereotaxic coordinates of the motor cortex are determined, and the skull is pierced at this location. A cannula prefilled with an AAV expressing cofilinS3D, an inactive form of cofilin, is slowly positioned in the cortical tissue. After AAV infusion, gold-covered screws (ECoG electrodes) are screwed through the skull and cemented to the skull with gold wires inserted in the neck muscles (EMG electrodes). The animals are allowed three weeks to recover and to ensure sufficient expression of cofilinS3D. The infected area and cell type are verified using immunohistochemistry, and the ECoG is analyzed using visual identification of vigilance states and spectral analysis. In summary, this combined methodological approach allows the investigation of the precise contribution of molecular components regulating neuronal morphology and connectivity to the regulation of synchronized cerebral cortex activity during wakefulness and sleep.

This article describes a protocol for the manipulation of molecular targets in the cerebral cortex using adeno-associated viruses and for monitoring the effects of this manipulation during wakefulness and sleep using electrocorticographic recordings.

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. ...