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>

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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><tbody><tr><td>C57BL/6J-Tg(Thy1-GCaMP6s)GP4.3Dkim/J</td><td>Jackson</td><td>24275</td><td>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><td>B6.Cg-Tg(Thy1-EGFP)OJrs/GfngJ.</td><td>Jackson</td><td>7919</td><td>Can be any strains expressing EGFP in subsets of neurons within specific populations in the cortex; providing a bright, vital Golgi-like stain.&amp;nbsp;</td></tr><tr><td>Dumont no. 5 forceps, standard, Dumoxel</td><td>Fine Science Tools</td><td>11251-30</td><td>Can be any brand of choice.</td></tr><tr><td>Dumont no. 5SF forceps</td><td>Fine Science Tools</td><td>11252-00</td><td>Can be any brand of choice.</td></tr><tr><td>Micro Bead Sterilizer</td><td>Southern Labware</td><td>B1201</td><td>Can be any brand of choice.</td></tr><tr><td>Harishige&amp;rsquo;s SG-4N Head Holding Adaptor</td><td>Tritech Research</td><td>SG-4N</td><td>Can be any brand of choice.</td></tr><tr><td>Ideal Micro-Drill Complete Kit</td><td>Harvard Apparatus</td><td>72-6065</td><td>Can be any brand of choice.</td></tr><tr><td>Burrs for Micro Drill</td><td>Fine Science Tools</td><td>19007-05</td><td>Can be any brand of choice.</td></tr><tr><td>Round cover glass, 3 mm</td><td>Harvard Biosciences</td><td>W4 64-0720</td><td>Can be any brand of choice.</td></tr><tr><td>Round cover glass, 5 mm</td><td>Harvard Biosciences</td><td>W4 64-0700</td><td>Can be any brand of choice.</td></tr><tr><td>Norland optical adhesive</td><td>Norland Products</td><td>7106</td><td>Can be any brand of choice.</td></tr><tr><td>Loctite liquid super glue</td><td>WB Mason</td><td>LOC1647358</td><td>Can be any brand of choice.</td></tr><tr><td>Grip cement kit, powder and solvent</td><td>Dentsply</td><td>675570</td><td>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><td>Gel foam</td><td>Moore Medical</td><td>2928</td><td>Can be any brand of choice.</td></tr><tr><td>Micro dissecting scissors</td><td>Roboz</td><td>RS-5841</td><td>Can be any brand of choice.</td></tr><tr><td>Artificial cerebrospinal fluid (ASF)</td><td>-</td><td>-</td><td>The ACSF contains 125 mM NaCl, 4.5 mM KCl, 26 mM NaHCO&lt;sub&gt;3&lt;/sub&gt;, 1.25 mM NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 2 mM CaCl&lt;sub&gt;2&lt;/sub&gt;, 1 mM MgCl&lt;sub&gt;2&lt;/sub&gt; 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 ...

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Neuroscience

11.0K Views.  Indiana University School of Medicine. Deze methode maakt het mogelijk de neurale activiteit gedurende een langere periode in vivo vast te registreren en te kwantificeren in bilaterale corticale regio's van de muis. De twee foton techniek heeft het voordeel van het detecteren van activiteit tegelijkertijd in grote of verspreide neuronale bevolking over een langere periode. De implicaties van deze techniek uitgebreid naar het onderzoek van activiteitspatronen in gedefinieerde neuronale populaties in verschillende regio's van de lev...

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

Flat-floored Air-lifted Platform: A New Method for Combining Behavior with Microscopy or Electrophysiology on Awake Freely Moving Rodents

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a living animal&#8217;s brain. Moreover, the lengthy recovery from anesthesia limits the frequency of repeated recording/imaging episodes in longitudinal studies. Hence, new methods that would allow stable recordings from non-anesthetized behaving mice are expected to advance the fields of cellular and cognitive neurosciences. Existing solutions range from mere physical restraint to more sophisticated approaches, such as linear and spherical treadmills used in combination with computer-generated virtual reality. Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage and explore its environment under stress-free conditions. This method allows researchers to perform behavioral tests (e.g., learning, habituation or novel object recognition) simultaneously with two-photon microscopic imaging and/or patch-clamp recordings, all combined in a single experiment. This video-article describes the use of the awake animal head fixation device (mobile homecage), demonstrates the procedures of animal habituation, and exemplifies a number of possible applications of the method.

This method creates a tangible, familiar environment for the mouse to navigate and explore during microscopic imaging or single-cell electrophysiological recordings, which require firm fixation of the animal&#8217;s head.

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a ...

Behavior

Multi-layer Cortical Ca2+ Imaging in Freely Moving Mice with Prism Probes and Miniaturized Fluorescence Microscopy

In vivo circuit and cellular level functional imaging is a critical tool for understanding the brain in action. High resolution imaging of mouse cortical neurons with two-photon microscopy has provided unique insights into cortical structure, function and plasticity. However, these studies are limited to head fixed animals, greatly reducing the behavioral complexity available for study. In this paper, we describe a procedure for performing chronic fluorescence microscopy with cellular-resolution across multiple cortical layers in freely behaving mice. We used an integrated miniaturized fluorescence microscope paired with an implanted prism probe to simultaneously visualize and record the calcium dynamics of hundreds of neurons across multiple layers of the somatosensory cortex as the mouse engaged in a novel object exploration task, over several days. This technique can be adapted to other brain regions in different animal species for other behavioral paradigms.

Multi-layer Cortical Ca2+ Imaging in Freely Moving Mice with Prism Probes and Miniaturized Fluorescence Microscopy

Here, we present a procedure for performing large-scale Ca2+ imaging with cellular-resolution across multiple cortical layers in freely moving mice. Hundreds of active cells can be observed simultaneously using a miniature, head-mounted microscope coupled with an implanted prism probe.

In vivo circuit and cellular level functional imaging is a critical tool for understanding the brain in action. High resolution imaging of mouse cortical ...

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits&#160;in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.

We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations&#160;in vitro.

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the ...

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

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

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

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

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

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in the brain sense various biological conditions in the periphery and transmit the signals to neurons. Microglia, immune cells in the brain, are involved in synaptic development and plasticity. Therefore, the contribution of microglia to neural circuit construction in response to the internal state of the body should be tested critically by intravital imaging of the relationship between microglial dynamics and neuronal activity.
Here, we describe a technique for the simultaneous imaging of microglial dynamics and neuronal activity in awake mice. Adeno-associated virus encoding R-CaMP, a gene-encoded calcium indicator of red fluorescence protein, was injected into layer 2/3 of the primary visual cortex in CX3CR1-EGFP transgenic mice expressing EGFP in microglia. After viral injection, a cranial window was installed onto the brain surface of the injected region. In vivo two-photon imaging in awake mice 4 weeks after the surgery demonstrated that neural activity and microglial dynamics could be recorded simultaneously at the sub-second temporal resolution. This technique can uncover the coordination between microglial dynamics and neuronal activity, with the former responding to peripheral immunological states and the latter encoding the internal brain states.

Here, we describe a protocol combining adeno-associated virus injection with cranial window implantation for simultaneous imaging of microglial dynamics and neuronal activity in awake mice.

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in ...

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the cytoplasm or the release of intracellular Ca2+ stores. Pirt-GCaMP3-based Ca2+ imaging of primary sensory neurons of the dorsal root ganglion (DRG) in mice offers the advantage of simultaneous measurement of a large number of cells. Up to 1,800 neurons can be monitored, allowing neuronal networks and somatosensory processes to be studied as an ensemble in their normal physiological context at a populational level in vivo. The large number of neurons monitored allows the detection of activity patterns that would be challenging to detect using other methods. Stimuli can be applied to the mouse hindpaw, allowing the direct effects of stimuli on the DRG neuron ensemble to be studied. The number of neurons producing Ca2+ transients as well as the amplitude of Ca2+ transients indicates sensitivity to specific sensory modalities. The diameter of neurons provides evidence of activated fiber types (non-noxious mechano vs. noxious pain fibers, A&#946;, A&#948;, and C fibers). Neurons expressing specific receptors can be genetically labeled with td-Tomato and specific Cre recombinases together with Pirt-GCaMP. Therefore, Pirt-GCaMP3 Ca2+ imaging of DRG provides a powerful tool and model for the analysis of specific sensory modalities and neuron subtypes acting as an ensemble at the populational level to study pain, itch, touch, and other somatosensory signals.

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

This protocol describes the surgical exposure of the dorsal root ganglion (DRG) followed by GCaMP3 (genetically-encoded Ca2+ indicator; Green Fluorescent Protein-Calmodulin-M13 Protein 3) Ca2+ imaging of the neuronal ensembles using Pirt-GCaMP3 mice while applying a variety of stimuli to the ipsilateral hind paw.

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the ...

Neuronal Activity Mapping in Developing Mouse Sensory Cortex

In Vivo Visualization of Spontaneous Activity in Neonatal Mouse Sensory Cortex at a Single-Neuron Resolution

The mammalian brain undergoes dynamic developmental changes at both the cellular and circuit levels throughout prenatal and postnatal periods. Following the discovery of numerous genes contributing to these developmental changes, it is now known that neuronal activity also substantially modulates these processes. In the developing cerebral cortex, neurons exhibit synchronized activity patterns that are specialized to each primary sensory area. These patterns markedly differ from those observed in the mature cortex, emphasizing their role in regulating area-specific developmental processes. Deficiencies in neuronal activity during development can lead to various brain diseases. These findings highlight the need to examine the regulatory mechanisms underlying activity patterns in neuronal development. This paper summarizes a series of protocols to visualize primary sensory areas and neuronal activity in neonatal mice, to image the activity of individual neurons within the cortical subfields using two-photon microscopy in vivo, and to analyze subfield-related activity correlations. We show representative results of patchwork-like synchronous activity within individual barrels in the somatosensory cortex. We also discuss various potential applications and some limitations of this protocol.

Author Spotlight: Deciphering Neural Circuit Formation from Two-Photon Microscopy and Single Neuron Imaging

Primary sensory areas in the neocortex exhibit unique spontaneous activities during development. This article describes how to visualize individual neuron activities and primary sensory areas to analyze area-specific synchronous activities in neonatal mice in vivo.

The mammalian brain undergoes dynamic developmental changes at both the cellular and circuit levels throughout prenatal and postnatal periods. Following ...

Advanced Imaging of Dorsal Root Ganglion Neurons for Sensory Modality Research

In Vivo Calcium Imaging of Dorsal Root Ganglia Neurons' Response to Somatic and Visceral Stimuli

A technique is described for surgically exposing the dorsal root ganglion (DRG) of the lumbar-6 in a live, anesthetized laboratory mouse, along with the protocol for in vivo calcium imaging of the exposed DRG in response to various visceral and somatic stimuli. Pirt-GCaMP6s mice or C57BL6 mice intrathecally injected with AAV viruses packaged with GCaMP6s were utilized to capture Ca2+ transients. The amplitude of these transients indicates sensitivity to specific sensory modalities. Afferent fibers originate from internal organs, with primary neuronal cell bodies in spinal or vagal ganglia. Studies on visceral nociception and acupuncture analgesia can potentially be conducted on primary sensory neurons using advanced imaging technologies like in vivo calcium imaging, allowing for the recording of neuronal activity ensembles in the intact animal during stimulation or intervention. The responses of DRG neuron ensembles to somatic and visceral stimuli applied to their corresponding receptive fields were recorded. This technique illustrates how neuronal populations react to various types of somatic and visceral stimuli. It is possible to comprehensively compare neuronal ensemble responses to different stimuli, which is a particularly valuable approach in research on visceral pain and segmental mechanisms of somatic stimulation, such as acupuncture.

The present protocol outlines in vivo calcium imaging for measuring the responses of ensembles of lumbar-6 DRG neurons to somatic and visceral stimuli. Thorough comparisons can be made among neurons responding to different stimuli. This protocol is valuable for investigating mechanisms of visceral pain and somatic stimulation, such as acupuncture.

A technique is described for surgically exposing the dorsal root ganglion (DRG) of the lumbar-6 in a live, anesthetized laboratory mouse, along with the ...

Calcium Imaging of Thoracic Segment T1 DRG for Cardiopulmonary Sensory Neurons

In Vivo Thoracic Dorsal Root Ganglia (DRG) Calcium Imaging and ECG Recording for Studying Peripheral Nerve Stimulation

The dorsal root ganglia (DRG), housing primary sensory neurons, transmit somatosensory and visceral afferent inputs to the dorsal horn of the spinal cord. They play a pivotal role in both physiological and pathological states, including neuropathic and visceral pain. In vivo calcium imaging of DRG enables real-time observation of calcium transients in single units or neuron ensembles. Accumulating evidence indicates that DRG neuronal activities induced by somatic stimulation significantly affect autonomic and visceral functions. While lumbar DRG calcium imaging has been extensively studied, thoracic segment DRG calcium imaging has been less explored due to surgical exposure and stereotaxic fixation challenges. Here, we utilized in vivo calcium imaging at the thoracic1 dorsal root ganglion (T1-DRG) to investigate changes in neuronal activity resulting from somatic stimulations of the forelimb. This approach is crucial for understanding the somato-cardiac reflex triggered by peripheral nerve stimulations (PENS), such as acupuncture. Notably, synchronization of cardiac function was observed and measured by electrocardiogram (ECG), with T-DRG neuronal activities, potentially establishing a novel paradigm for somato-visceral reflex in the thoracic segments.

This study presents a surgical manipulation to expose the T-DRG in anesthetized mice for in vivo calcium imaging, along with synchronous ECG recordings. This method represents a cutting-edge tool for studying peripheral electric nerve stimulation and thoracic visceral organ inputs, as well as their interactions at the primary sensory level.

The dorsal root ganglia (DRG), housing primary sensory neurons, transmit somatosensory and visceral afferent inputs to the dorsal horn of the spinal cord. ...

Installation of Optical Windows in the Brain of Transgenic Mice to Image Neural Activities

Source: Lin, X., et al. <a href="https://app.jove.com/v/56297">Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice.</a> J. Vis. Exp. (2019).
This video demonstrates the brain&#39;s neural activity recording through an optical window in transgenic mice. The mice express a fluorescent protein that enables the detection of neural activities. A small portion of the skull is carefully removed to install an optical window, and the skull is then covered with black cement to block light before imaging.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

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