installation of optical windows in the brain of transgenic mice to image neural activities

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

Confirm a lack of response to toe pinch on an anesthetized 20 to 25 gram, 1 and 1/2 to 2 and 1/2-month-old Thy1-GCaMP6s mouse and apply ointment to the animal&#39;s eyes. Place the mouse on a heating pad covered with a sterile drape, and use a head-holding adapter for mice to stabilize the head. Shave most of the scalp with a double-edged razor, and clean the exposed skin with sequential sterile alcohol preparation pads and povidone-iodine solution scrubs. Then, use a pair of scissors to make a rectangular 2-by-3-millimeter cut in the scalp.
Using a cotton swab, push the skin aside to create an exposure area of greater than 3 millimeters in diameter, and use a blunt microsurgical blade to gently remove the connective tissue attached to the skull. With a dental drill, gently mark a 3-millimeter diameter circle around the S1 area. After making a similar circle on the left side of the skull, apply a thin layer of cyanoacrylate superglue to both sides of the bone to provide a base for dental cement application.
Under a dissecting microscope, use a high-speed micro-drill to thin down a circular groove in the right side of the skull around the S1 area to create a smooth edge, aspirating the bone debris with a vacuum as necessary. After an approximately 2/3rd bone depth has been achieved, slowly and carefully thin the remaining one-third of bone until a circular bone flap has been completely freed from the surrounding skull. Use a pair of # 5/45 forceps to slowly and carefully remove the circular piece of bone to expose the dura, taking care to avoid damaging the pial vessels.
After removing a bone flap from the left side of the skull in the same manner, rinse the optical window for the right side of the skull with sterile saline and check for imperfections under a stereomicroscope. Install the optical window over the craniotomy region with the top portion of the window resting on the skull and the bottom portion within the craniotomized opening, resting on the dura in the presence of cerebral spinal fluid. Use the cyanoacrylate super glue to seal the top portion of the optical window edge to the skull.
When the glue has dried, apply black dental cement to the edge of the glass, the rest of the right side of the exposed skull, and the wound margins to block the light. Then, install the left optical window and allow the animal to recover with monitoring until full recumbency. 7 to 10 days after the procedure, place the anesthetized animal into the head-holding adapter on a heating pad, under a two-photon microscope, taking care that the optical window and the right side of the skull are oriented perpendicular to the optical axis of the microscope.
Acquire an initial reference map image of the cranial window using brightfield illumination at the 4 times magnification. After acquiring multiple images of the same region of interest, image the calcium dynamics within the S1 area in the left hemisphere and calculate the changes in the fluorescence from each region of interest on both sides of the skull.

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Nanyang Technological University

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Imaging and Optical Techniques

EoE Sub Articles

eoe sub articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Preparing the Animal for Imaging
<ol>
	<li>Place sterile gauze, cotton swabs, and autoclaved surgical tools in the surgery area. 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 of 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 a 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 1A). Remove the connective tissue attached to the skull by gently scraping the skull with a blunt microsurgical blade (Scalpel Blade #10; Figure 1C).</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 1D). The purpose is to create a smooth edge for the 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 1E).</li>
	<li>Keep the exposed brain region moist with ACSF (Figure 1E). 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 1F).</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 1B).</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>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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.</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&#39;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 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>

Source: Lin, X., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/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.

<img alt="Figure 1" src="/files/ftp_upload/22749/22749_Figure1.jpg" /> 
Figure 1. Creation of bilateral windows on the skull. (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 the 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.
<img alt="Figure 2" src="/files/ftp_upload/22749/22749_Figure2.jpg" /> 
Figure 2: Schematic drawing shows key steps for preparing a cranial window. (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.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Begin with an anesthetized, genetically altered mouse with its head secured in a head-holding adaptor.
The mouse brain expresses a fluorescent protein that enables the detection of neural activities.
Shave and sequentially sterilize the scalp before cutting it open.
Remove connective tissue to expose the skull.
Mark the target areas and apply a layer of glue.
Use a microdrill to thin the skull on the marked area.
Remove the piece of the skull to expose the brain.
Apply artificial cerebrospinal fluid to keep the brain moist.
Install an optical window in the area and seal it with glue, which provides a base for the cement attachment.
Apply black dental cement to block the light.
Repeat the procedure on the other side of the skull.
Allow the animal to recover.
To image neural activity, place the secured anesthetized mouse under the microscope and acquire images.

53 Views. تركيب النوافذ البصرية في دماغ الفئران المعدلة وراثيا لتصوير الأنشطة العصبية

Video: تركيب النوافذ البصرية في دماغ الفئران المعدلة وراثيا لتصوير الأنشطة العصبية - Experiment

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.

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.

التصوير النشاط العصبي في القشرة الأولية Somatosensory باستخدام Thy1-GCaMP6s الفئران المحورة وراثيا

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JoVE Journal

Visualizing Protein Kinase A Activity In Head-fixed Behaving Mice Using In Vivo Two-photon Fluorescence Lifetime Imaging Microscopy

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. Despite their importance, technologies for tracking neuromodulatory events with cellular resolution are just beginning to emerge. Neuromodulators, such as dopamine, norepinephrine, acetylcholine, and serotonin, trigger intracellular signaling events via their respective G protein-coupled receptors to modulate neuronal excitability, synaptic communications, and other neuronal functions, thereby regulating information processing in the neuronal network. The above mentioned neuromodulators converge onto the cAMP/protein kinase A (PKA) pathway. Therefore, in vivo PKA imaging with single-cell resolution was developed as a readout for neuromodulatory events in a manner analogous to calcium imaging for neuronal electrical activities. Herein, a method is presented to visualize PKA activity at the level of individual neurons in the cortex of head-fixed behaving mice. To do so, an improved A-kinase activity reporter (AKAR), called tAKAR&#945;, is used, which is based on F&#246;rster resonance energy transfer (FRET). This genetically-encoded PKA sensor is introduced into the motor cortex via in utero electroporation (IUE) of DNA plasmids, or stereotaxic injection of adeno-associated virus (AAV). FRET changes are imaged using two-photon fluorescence lifetime imaging microscopy (2pFLIM), which offers advantages over ratiometric FRET measurements for quantifying FRET signal in light-scattering brain tissue. To study PKA activities during enforced locomotion, tAKAR&#945; is imaged through a chronic cranial window above the cortex of awake, head-fixed mice, which run or rest on a speed-controlled motorized treadmill. This imaging approach will be applicable to many other brain regions to study corresponding behavior-induced PKA activities and to other FLIM-based sensors for in vivo imaging.

A procedure is presented to visualize protein kinase A activities in head-fixed, behaving mice. An improved A-kinase activity reporter, tAKAR&#945;, is expressed in cortical neurons and made accessible for imaging through a cranial window. Two-photon fluorescence lifetime imaging microscopy is used to visualize PKA activities in vivo during enforced locomotion.

تصور البروتين كيناز نشاط في الفئران يتصرف الرأس ثابتة باستخدام في فيفو اثنين من الفوتون الفلورة التصوير مدى الحياة المجهرية

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. ...

Multi-Fiber Photometry to Record Neural Activity in Freely-Moving Animals

Recording the activity of a group of neurons in a freely-moving animal is a challenging undertaking. Moreover, as the brain is dissected into smaller and smaller functional subgroups, it becomes paramount to record from projections and/or genetically-defined subpopulations of neurons. Fiber photometry is an accessible and powerful approach that can overcome these challenges. By combining optical and genetic methodologies, neural activity can be measured in deep brain structures by expressing genetically-encoded calcium indicators, which translate neural activity into an optical signal that can be easily measured. The current protocol details the components of a multi-fiber photometry system, how to access deep brain structures to deliver and collect light, a method to account for motion artifacts, and how to process and analyze fluorescent signals. The protocol details experimental considerations when performing single and dual color imaging, from either single or multiple implanted optic fibers.

This protocol details how to implement and perform multi-fiber photometry recordings, how to correct for calcium-independent artifacts, and important considerations for dual-color photometry imaging.

القياس الضوئي متعدد ألياف لتسجيل النشاط العصبي في الكائنات المتحركة بحريه

Recording the activity of a group of neurons in a freely-moving animal is a challenging undertaking. Moreover, as the brain is dissected into smaller and ...

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

Transcript

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