This work was supported by a Natural Sciences and Engineering Research Council of Canada (NSERC) Discovery Grant #40352, Campus Alberta for Innovation Program Chair, Alberta Alzheimer Research Program to MHM, and NSERC CREATE in BIF doctoral fellowship and AIHS postgraduate fellowship to MK. We thank Pu Min Wang for the development of this protocol and for surgical training, and Behroo Mirza Agha and Di Shao for husbandry.

The following protocol follows the University of Lethbridge Animal Care Committee (ACC) guidelines, and is conducted in accordance with the standards of the Canadian Council on Animal Care (CCAC).
1. Preparation
<ol>
	<li>For prolonged study periods, autoclave all opened surgical supplies and ensure that sterility is maintained throughout the surgery. If multiple surgeries are required, autoclave between surgeries.&#160;</li>
	<li>Ensure there is plenty of brain buffer on hand (at least 50 mL). The solution is comprised of 134 mM sodium chloride, 5.4 mM potassium, 1 mM magnesium chloride hexahydrate, 1.8 mM calcium chloride dihydrate, and 5 mM HEPES sodium, pH balanced to 7.4 with 5 M hydrogen chloride.</li>
	<li>Place the mouse in an induction chamber and anesthetize with 3 - 4% isoflurane. Follow with 1.0 - 2.0% isoflurane for maintenance during the surgery.&#160;Further reduce to as low as 0.4 &#8211; 0.8% during imaging5,13,14,15, provided proper anesthesia is maintained. These low levels of anesthesia require vigilant and frequent monitoring of the mouse (about every 5 min) to ensure it remains areflexic to painful stimuli. 
	NOTE:&#160;Dehydration can become problematic due to the high surface to volume ratio of the mouse and exacerbated by prolonged use of isoflurane.
	<ol>
		<li>Use subcutaneous injections of saline, 0.1 mL per 10 g body weight, every 1 - 2 h. When adequately hydrated, the mouse will urinate once every 1 - 2 h.</li>
	</ol>
	</li>
	<li>Closely monitor the mouse to ensure consistent anesthesia throughout surgery and imaging. Do not leave the mouse unattended and take care that it never regains consciousness.</li>
	<li>Transfer the anesthetized mouse to the head-holder set-up and place on a thermo-regulating heating pad set to 37 &#176;C. Secure the upper teeth in a teeth holder.</li>
	<li>Rotate the mouse&#39;s head towards the left approximately 30&#176; to expose the right lateral side of the head and secure the mouse&#39;s head with the blunt end of ear bars (Figure 1A).</li>
	<li>Apply ophthalmic ointment to prevent corneal drying.</li>
	<li>To reduce cerebral edema, inject dexamethasone (4 mg/kg) intramuscularly.</li>
	<li>Wipe the skin over the surgical area with cotton swabs dipped in 4% chlorhexidine (3 times) and 70% ethanol (3 times). Use each cotton swab only once.</li>
	<li>Don surgical gloves and cover the animal with adhesive plastic wrap. Provide local anesthetic by injecting of lidocaine (8 - 10 mg/kg; 2% epinephrine) subcutaneously over the craniotomy site. Wait 3 - 5 min for the drug to be absorbed into the tissue.</li>
</ol>
2. Removing the Skin and Retracting the Muscle from the Skull
<ol>
	<li>Perform nearly all of these procedures while viewing the skull under a dissecting microscope (e.g., 0.7 - 4.5X power, depending on the situation).</li>
	<li>Lift up the skin 1 mm left of the midline (just behind the ear) with forceps and make a small horizontal incision with surgical scissors.</li>
	<li>Make a 5 - 6 mm lateral cut towards the right ear, and then cut towards the rostral end of the head.</li>
	<li>At the initial incision point, insert the scissors and cut 10 mm rostrally.</li>
	<li>Cut the skin around the right ear and near the right eye to expose the right side of the skull and temporal muscle. Ensure that the widest part of the exposed area is at least 7 mm. Trim the skin further if the surgical area needs to be extended.</li>
	<li>Fix the skin around the incision by putting a few drops of butyl cyanoacrylate glue between the skull and the skin.&#160;Ensure the tissue has been previously dried with cotton tipped applicators or rolled tissues to increase the efficacy of the adhesive.</li>
	<li>Using a cotton swab, rub the surface of the skull in a circular motion to remove the periosteum from the skull. Ensure that none remains by drying the skull completely.</li>
	<li>Using spring scissors and forceps, separate the temporal muscle from the skull; cut and retract the muscle laterally until reaching the squamosal bone (Figure 1C). Take extreme care not to damage the superficial temporal vein that runs along the level of the squamosal bone near the eye, otherwise hemorrhaging might occur.</li>
	<li>Control bleeding with gel foam pre-soaked in brain buffer. For serious hemorrhaging use a heat cauterizer.&#160;Drop butyl cyanoacrylate glue onto the bleeding sites following treatment with the heat cauterizer. Ensure the area is dried beforehand by using cotton tipped applicators or rolled tissues to increase the efficacy of the adhesive.&#160;</li>
</ol>
3. Craniotomy
NOTE: The surgeon must remain diligent during removal of the skull and dura to avoid unnecessary complications. Troubleshooting steps are included should complications arise.
<ol>
	<li>Mark the location of bregma either with a fine tip marker or cut a small triangular piece tape and point a corner at bregma (Figure 2A) to orient the underlying brain regions.</li>
	<li>Prior to affixing the head plate ensure the skull is completely dry. The skull will quickly air dry after application of the gel foam soaked in brain buffer, if more drying is needed, use cotton tip swabs. This step is crucial for head plate fixation (Figure 1B). 
	NOTE: If there is periosteum left on the skull, or if the skull is not dry before gluing on the head plate, it will likely detach. If this happens, gently remove the head plate and start over. Bleeding may occur during this process; allow a few minutes for it to clot, and then gently remove. This process is not recommended to be repeated more than twice.</li>
	<li>Apply ethyl cyanoacrylate glue around the bottom edge of the head plate16, and glue the head plate over the craniotomy area (Figure 1D &#38; Figure 2A).</li>
	<li>Fill the opening space between the skull and head plate with dental cement leaving only the craniotomy area exposed. Wait for the dental cement to dry and harden, typically 5 - 10 min (Figure 2B).
	<ol>
		<li>Once the cement is set, briefly fill the well with brain buffer and allow to soak for 3 - 5 min. Use a rolled tissue to remove brain buffer before drilling (Figure 2C).</li>
	</ol>
	</li>
	<li>Outline the surgical area by lightly scoring the surface of the skull with a dental drill. Use a pneumatic drill (set to maximum of 20 PSI), with a FG &#188; burr, and controlled with a variable speed foot pedal.</li>
	<li>Gently trace the drill along the original scoring to deepen it, ensuring the drill does not penetrate through the skull into the brain (Figure 2D). Take turns every few minutes between drilling and dabbing the skull surface with moistened rolled tissues. This will reduce heating and drying of the skull from mechanical friction and prolonged exposure. 
	NOTE: The skull will quickly air dry after application of the wet gel foam. If more drying is needed, use cotton tip swabs. 
	Caution: The skull is uneven in thickness. For example, the parietal-temporal ridge is the thickest area, while skull regions near the midline and squamosal landmarks are relatively thin.</li>
	<li>During drilling, periodically check for buckling of the skull by gently pressing on it with forceps or the non-moving drill bit. When the bone begins to buckle, stop drilling and immerse the entire window in brain buffer. 
	NOTE: If blood rushes out of an area, it may suggest that the dura has been damaged. If this is the case, place a semi-wet gel foam over the area and try to soak up the blood while gently applying pressure to the gel foam with a cotton tip swab.</li>
	<li>Wait for at least 5 min before skull removal to soften the bone and to reduce the chance of the dura sticking to the bone, making the skull removal process easier.</li>
	<li>Perform the skull removal process while the skull is submerged in brain buffer. 
	NOTE: If a portion of the skull remains stubbornly attached, a #11 scalpel blade can be used to gently score the skull. Take extreme care to not puncture the blade through the skull and into the brain.</li>
	<li>Beginning from the anterior edge, gently pry the loose skull from the dura using forceps. 
	NOTE: If a small amount of bleeding occurs during the skull removal process, remove the buffer with a transfer pipette or syringe, and then replace with new buffer.</li>
	<li>Once the bone is loose and &#34;floating&#34; on the dura, firmly grip the bone with forceps and lift the bone from the dura. Ensure the bone never penetrates into the brain.</li>
	<li>To control bleeding, roll the corner of a tissue into a point and remove most of the buffer from the cranial well. Quickly apply gel foam, pre-soaked in buffer, to the bleeding area while adding very light pressure with a cotton tip swab. 
	NOTE: The bleeding usually comes from the edge of the bone or the surface of the dura; both cases are normal and bleeding will quickly stop if no major blood vessels are damaged. If bleeding continues, blood may fill the entire window, forming a clot sheet over the imaging area.
	<ol>
		<li>To remove the clot sheet, carefully pick up pieces of clotted blood from the imaging area while leaving the blood clot intact around the source of the ruptured blood vessel. Take care to not remove the blood clot from the bleed source as this may cause even more blood loss. Irrigate the surface of the brain with brain buffer to wash away any blood.</li>
		<li>Take care to avoid touching the delicate brain tissue or adding foreign material to the brain; repeat until bleeding has stopped, for approximately 2 - 5 min (Figure 2E). 
		NOTE: At this point, the craniotomy is ready for preparing the cranial window (see step 5). If needed, remove the dura before implanting the cranial window (see step 4).</li>
	</ol>
	</li>
</ol>
4. Dura Removal
NOTE: Dura removal requires extreme care and may take over 15 min.
<ol>
	<li>Draw away excess buffer from the craniotomy. While maintaining a moist surface, grab a small piece of dura with forceps and gently tear the dura.</li>
	<li>Use forceps and spring scissors to gently tear and cut away the dura.</li>
	<li>Drop more brain buffer on the brain surface to float the dura and help it separate from the brain. Continue until all dura is removed from the cranial window site. When performed correctly the brain will appear to be very clean with distinct blood vessels and no blemishes (compare Figure 2E with 2F). 
	Caution: Some areas of the dura are attached to small arterioles on the surface of the brain (e.g., near the midline proximal to the parietal association area), and removal of such can rupture the arteriole. In such cases, it may be better to leave a small piece of dura intact over top of the arteriole. The VSD may not penetrate that small area, but this is preferred to having major bleeding.</li>
	<li>Fix the brain surface in agarose as soon as possible to minimize movement from pulsation and to prevent further swelling (see step 5).</li>
</ol>
5. Preparing Cranial Window
<ol>
	<li>Spray the cover glass with 70% ethanol and use an air canister to gently dry. Ensure the glass is completely clean with no spots or dust present.</li>
	<li>Prepare 1.3% agarose by heating 200 mg of agarose powder dissolved in 15 mL of brain buffer. Set the microwave to high and heat for 10 - 15 s at a time, gently stirring in between, until all agar is dissolved. 
	Note: Ensure no bubbles or particles are present, as these will interfere with imaging.</li>
	<li>Place a thermometer in the hot agarose and cool the agarose down to just above solidifying temperature (~ 40 &#176;C). 
	NOTE: Running cool water over the outside of the agarose container may speed the cooling process. Gently stir continuously to ensure no bubbles or particulates are present.</li>
	<li>Immediately before applying the agar, remove the brain buffer from the cranial well. Draw up the agarose with a transfer pipette and drop the agarose directly on the brain. Quickly place the cover slip over the surface and fasten the cover slip with agarose drops on the corners.</li>
</ol>
6. Euthanasia
NOTE: In our experience, this procedure takes experienced surgeons at least 3 - 4 practice surgeries to attain greater than 90% success rate. Less experienced surgeons may require even more practice. During the craniotomy or durotomy, the brain may sustain damage, such as if the drill punches through the bone into the brain. Some minor damage at the edge of craniotomy may be permissible. However, if the brain does not look &#34;clean&#34; with bright red undamaged blood vessels and white cortex, the experiment may need to be terminated. Examples of poor preparations include those with dead blood vessels, or when the cortex is marked with torn or damaged blood vessels. If any of these signs are present, the experiment will unlikely yield high quality data.&#160;Whether the surgery/experiment was successful or not, humanely euthanize the mouse at the completion of the experiment.
<ol>
	<li>Deeply anesthetize with at least 3.5% isoflurane. Then, give an intraperitoneal injection of sodium pentobarbital at 300 mg/kg. Ideally, if the needle is inserted into the liver, death will be very rapid (&#60; 1 min).</li>
	<li>If perfusion is required, verify that the animal is deeply anesthetized before proceeding (see step 1.3).</li>
	<li>Alternatively, if perfusion is not required, wait for a minimum of 5 min and then verify that the mouse is dead. Confirm the absence of respiration, heartbeat, as well as a lack of pain withdrawal and corneal reflexes. Also, observe for pale blue/white coloring of extremities and darkening of the blood vessels over the cortex.</li>
</ol>

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The authors have nothing to disclose.

This innovative protocol for a large cranial window enables simultaneous imaging over the temporal and parietal areas of the cerebral cortex. Combined with optical imaging, it can help to reveal neural dynamics within cortical areas during spontaneous and stimulus-induced activity. This expansive craniotomy also exposes a large extension of the cortical vasculature network, including the proximal end of the middle cerebral artery (MCA), enabling in vivo imaging of blood flow and direct manipulation of lateral ve...

The craniotomy is a standard procedure used by neuroscientists to reveal a portion of the brain. Since the dawn of electrophysiology, the craniotomy has allowed unprecedented breakthroughs in the field of neuroscience. Dense mapping of the cerebral cortex with electrodes has led to experiments testing hypotheses and theories based on these maps. We have recently entered a new era where the craniotomy is being utilized for in vivo imaging of cortical blood flow1,2,3 and neurovascular architecture4, enabling real time visualization of cortical activity within the exposed areas5,6,7. Although many studies use craniotomies combined with in vivo optical imaging techniques to study the structure and function of cortical neurons, glia, and cortical vasculature8,9, further investigations are limited by small areas of exposed cortex (but see10).
The purpose of this protocol is to provide a method for creating a large lateral craniotomy, exposing the cerebral cortex from the midline to the squamosal bone, and extending beyond bregma and lambda. This large craniotomy enables simultaneous viewing of the association cortices (retrosplenial, cingulate, and parietal), primary and secondary motor, somatosensory, visual, and the auditory cortex. This method has been previously coupled with voltage sensitive dye imaging (VSDI) to investigate how multiple cortical areas interact with one another during spontaneous and stimulus-induced cortical activity5,11,12. The most challenging aspects of this procedure include positioning the head of the animal, fixing the head plate, and avoiding hemorrhage while separating the temporal muscle from the parietal bone. Care must also be taken during the drilling and skull removal processes as the skull curves at an oblique angle.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Heating Pad&#160;</td>
			<td>FHC</td>
			<td>40-90-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors</td>
			<td>Fine Science Tools</td>
			<td>14058-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps&#160;</td>
			<td>Fine Science Tools</td>
			<td>11251-35</td>
			<td>2 or more pairs are recommended</td>
		</tr>
		<tr>
			<td>Spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-00, 15000-10</td>
			<td>1 pair should be designated for dura removal&#160;</td>
		</tr>
		<tr>
			<td>Jet tooth shade powder</td>
			<td>LANG Dental</td>
			<td>Jet Tooth Shade Powder</td>
			<td>to be mixed with the Jet Liquid</td>
		</tr>
		<tr>
			<td>Jet tooth shade liquid</td>
			<td>LANG Dental</td>
			<td>Jet Tooth Shade Liquid</td>
			<td>to be mixed wihth the Jet Powder&#160;</td>
		</tr>
		<tr>
			<td>Drill Heads - Carbide Burs FG 1/4 389</td>
			<td>Midwest Dental</td>
			<td>385201</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose Powder</td>
			<td>Sigma-Aldrich</td>
			<td>A9793</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelfoam</td>
			<td>Sinclair Dental Canada</td>
			<td>Pfizer Gelfoam</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Western Drug Distribution Centre Ltd</td>
			<td>124125</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine 2% Epinephrine</td>
			<td>Western Drug Distribution Centre Ltd</td>
			<td>125299</td>
			<td></td>
		</tr>
		<tr>
			<td>Dexamethazone 5 mg/mL</td>
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a large lateral craniotomy procedure for mesoscale wide-field optical imaging of brain activity

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, typically 3 mm x 3 mm. This protocol introduces a method for creating a substantially larger 7 mm x 6 mm cranial window exposing most of a cerebral hemisphere over the mouse temporal and parietal cortices (e.g., bregma 2.5 - 4.5 mm, lateral 0 - 6 mm). To perform this surgery, the head must be tilted approximately 30&#176; and much of the temporal muscle must be retracted. Due to the large amount of bone removal, this procedure is intended only for acute experiments with the animal anesthetized throughout the surgery and experiment.
The main advantage of this innovative large lateral cranial window is to provide simultaneous access to both medial and lateral areas of the cortex. This large unilateral cranial window can be used to study the neural dynamics between cells, as well as between different cortical areas by combining multi-electrode electrophysiological recordings, imaging of neuronal activity (e.g., intrinsic or extrinsic imaging), and optogenetic stimulation. Additionally, this large craniotomy also exposes a large area of cortical blood vessels, allowing for direct manipulation of the lateral cortical vasculature.

To study the interactions between cortical areas within a single hemisphere, we used a large craniotomy extending across the sagittal sinus and 5 - 6 mm lateral. This cranial window included primary (motor, somatosensory, visual, auditory), secondary (motor, visual), and association (retrosplenial, cingulate, parietal association) cortices of right brain hemispheres (Figure 3A). For this work we used voltage sensitive dye (VSD) imaging, which reflects changes...

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A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

This protocol presents a method for creating a large unilateral craniotomy over the temporal and parietal regions of the mouse cerebral cortex. This is especially useful for real time imaging over an expansive area of a cortical hemisphere.

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, ...

a-large-lateral-craniotomy-procedure-for-mesoscale-wide-field-optical

Research

JoVE Journal

Neuroscience

12.2K Views.  University of Lethbridge.  Dit protocol geeft een methode waarbij een grote unilaterale craniotomie via temporale en pariëtale gebieden van het muizen cerebrale cortex. Dit is vooral handig voor real-time beeldvorming over een uitgestrekt gebied van een corticale halfrond. 

Video: A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

State-Dependency Effects on TMS:  A Look at Motive Phosphene Behavior

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability (either increasing or decreasing it) via the application of localized magnetic field pulses.1,2 Within the field of TMS, the term state dependency refers to the initial, baseline condition of the particular neural region targeted for stimulation. As can be inferred, the effects of TMS can (and do) vary according to this primary susceptibility and responsiveness of the targeted cortical area.3,4,5

In this experiment, we will examine this concept of state dependency through the elicitation and subjective experience of motive phosphenes. Phosphenes are visually perceived flashes of small lights triggered by electromagnetic pulses to the visual cortex. These small lights can assume varied characteristics depending upon which type of visual cortex is being stimulated. In this particular study, we will be targeting motive phosphenes as elicited through the stimulation of V1/V2 and the V5/MT+ complex visual regions.6

In this article, we examine the effects of visually relevant state dependency on TMS induced motive phosphenic presentations.

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate ...

Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various neurological deficits. The current ex vivo live imaging of motile ependymal cilia technique allows for a detailed study of ciliary dynamics following several steps. These steps include: mice euthanasia with carbon dioxide according to protocols of The University of Toledo&#8217;s Institutional Animal Care and Use Committee (IACUC); craniectomy followed by brain removal and sagittal brain dissection with a vibratome or sharp blade to obtain very thin sections through the brain lateral ventricles, where the ependymal cilia can be visualized. Incubation of the brain&#8217;s slices in a customized glass-bottom plate containing Dulbecco&#8217;s Modified Eagle&#8217;s Medium (DMEM)/High-Glucose at 37 &#176;C in the presence of 95%/5% O2/CO2 mixture is essential to keep the tissue alive during the experiment. A video of the cilia beating is then recorded using a high-resolution differential interference contrast microscope. The video is then analyzed frame by frame to calculate the ciliary beating frequency. This allows distinct classification of the ependymal cells into three categories or types based on their ciliary beating frequency and angle. Furthermore, this technique allows the use of high-speed fluorescence imaging analysis to characterize the unique intracellular calcium oscillation properties of ependymal cells as well as the effect of pharmacological agents on the calcium oscillations and the ciliary beating frequency. In addition, this technique is suitable for immunofluorescence imaging for ciliary structure and ciliary protein localization studies. This is particularly important in disease diagnosis and phenotype studies. The main limitation of the technique is attributed to the decrease in live motile cilia movement as the brain tissue starts to die.

Using high-resolution differential interference contrast (DIC) microscopy, an ex vivo observation of the beating of motile ependymal cilia located within the mouse brain ventricles is demonstrated by live-imaging. The technique allows a recording of the unique ciliary beating frequency and beating angle as well as their intracellular calcium oscillation pacing properties.

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various ...

Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study these two systems and how they interact has been with the use of in vivo preparations. Major advances have been made with acute brain slices containing the thalamocortical and cortico-thalamic pathways in the somatosensory, visual, and auditory systems. With key refinements to our recent modification of the auditory thalamocortical slice1, we are able to more reliably capture the projections between most of the major auditory midbrain and forebrain structures: the inferior colliculus (IC), medial geniculate body (MGB), thalamic reticular nucleus (TRN), and the auditory cortex (AC). With portions of all these connections retained, we are able to answer detailed questions that complement the questions that can be answered with in vivo preparations. The use of flavoprotein autofluorescence imaging enables us to rapidly assess connectivity in any given slice and guide the ensuing experiment. Using this slice in conjunction with recording and imaging techniques, we are now better equipped to understand how information processing occurs at each point in the auditory forebrain as information ascends to the cortex, and the impact of descending cortical modulation. 3-D printing to build slice chamber components permits double-sided perfusion and broad access to networks within the slice and maintains the widespread connections key to fully utilizing this preparation.

Using multiple angles to cut the mouse pup brain, we improve upon a previously-described acute brain slice which captures the connections between most of the major auditory midbrain and forebrain structures.

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study ...

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.

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

Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical properties, synaptic connections, and in vivo functions, but their basic functional and anatomical organization from cellular to network scale is poorly understood. Here we describe a method for the three-dimensional imaging of fluorescently-labeled neurons across large areas of the brain for the investigation of the cortical cellular organization. Specific types of neurons are labeled by the injection of fluorescent retrograde neuronal tracers or expression of fluorescent proteins in transgenic mice. Block brain samples, e.g., a hemisphere, are prepared after fixation, made transparent with tissue clearing methods, and subjected to fluorescent immunolabeling of the specific cell types. Large areas are scanned using confocal or two-photon microscopes equipped with large working distance objectives and motorized stages. This method can resolve the periodic organization of the cell type-specific microcolumn functional modules in the mouse neocortex. The procedure can be useful for the study of three-dimensional cellular architecture in the diverse brain areas and other complex tissues.

Here we describe a procedure for tissue clearing, fluorescent labeling, and large-scale imaging of mouse brain tissue which, thereby, enables visualization of the three-dimensional organization of cell types in the neocortex.

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Real-Time fMRI Brain Mapping in Animals

Dynamic fMRI responses vary largely according to the physiological conditions of animals either under anesthesia or in awake states. We developed a real-time fMRI platform to guide experimenters to monitor fMRI responses instantaneously during acquisition, which can be used to modify the physiology of animals to achieve desired hemodynamic responses in animal brains. The real-time fMRI set-up is based on a 14.1T preclinical MRI system, enabling the real-time mapping of dynamic fMRI responses in the primary forepaw somatosensory cortex (FP-S1) of anesthetized rats. Instead of a retrospective analysis to investigate confounding sources leading to the variability of fMRI signals, the real-time fMRI platform provides a more effective scheme to identify dynamic fMRI responses using customized macro-functions and a common neuroimage analysis software in the MRI system. Also, it provides immediate troubleshooting feasibility and a real-time biofeedback stimulation paradigm for brain functional studies in animals.

Animal brain functional mapping can benefit from the real-time functional magnetic resonance imaging (fMRI) experimental set-up. Using the latest software implemented in the animal MRI system, we established a real-time monitoring platform for small animal fMRI.

Dynamic fMRI responses vary largely according to the physiological conditions of animals either under anesthesia or in awake states. We developed a ...

Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates reafferent cues that animals must disentangle from relevant sensory cues of the surrounding environment. For example, when a fish swims, flow generated from body undulations is detected by the mechanoreceptive neuromasts, comprising hair cells, that compose the lateral line system. The hair cells then transmit fluid motion information from the sensor to the brain via the sensory afferent neurons. Concurrently, corollary discharge of the motor command is relayed to hair cells to prevent sensory overload. Accounting for the inhibitory effect of predictive motor signals during locomotion is, therefore, critical when evaluating the sensitivity of the lateral line system. We have developed an in vivo electrophysiological approach to simultaneously monitor posterior lateral line afferent neuron and ventral motor root activity in zebrafish larvae (4-7 days post fertilization) that can last for several hours. Extracellular recordings of afferent neurons are achieved using the loose patch clamp technique, which can detect activity from single or multiple neurons. Ventral root recordings are performed through the skin with glass electrodes to detect motor neuron activity. Our experimental protocol provides the potential to monitor endogenous or evoked changes in sensory input across motor behaviors in an intact, behaving vertebrate.

We describe a protocol to monitor changes in the afferent neuron activity during motor commands in a model vertebrate hair cell system.

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates ...

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit functions in systems neuroscience. However, high absorbance and scattering of light in mammalian tissue limit intravital imaging mostly to superficial brain regions, leaving deep-brain areas, such as the hippocampus, out of reach for optical microscopy. In this video, we show the preparation and implantation of the custom-made imaging window to enable chronic in vivo imaging of the dorsal hippocampal CA1 region in head-fixed behaving mice. The custom-made window is supplemented with an infusion cannula that allows targeted delivery of viral vectors and drugs to the imaging area. By combining this preparation with wide-field imaging, we performed a long-term recording of neuronal activity using a fluorescent calcium indicator from large subsets of neurons in behaving mice over several weeks. We also demonstrated the applicability of this preparation for voltage imaging with single-spike resolution. High-performance genetically encoded indicators of neuronal activity and scientific CMOS cameras allowed the recurrent visualization of subcellular morphological details of single neurons at high temporal resolution. We also discuss the advantages and potential limitations of the described method and its compatibility with other imaging techniques.

This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice.

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit ...

In Vivo Wide-Field and Two-Photon Calcium Imaging from a Mouse Using a Large Cranial Window

Wide-field calcium imaging from the mouse&#39;s neocortex allows one to observe cortex-wide neural activity related to various brain functions. On the other hand, two-photon imaging can resolve the activity of local neural circuits at the single-cell level. It is critical to make a large cranial window to perform multiple-scale analysis using both imaging techniques in the same mouse. To achieve this, one must remove a large section of the skull and cover the exposed cortical surface with transparent materials. Previously, glass skulls and polymer-based cranial windows have been developed for this purpose, but these materials are not easily fabricated. The present protocol describes a simple method for making a large cranial window consisting of commercially available polyvinylidene chloride (PVDC) wrapping film, a transparent silicone plug, and a cover glass. For imaging the dorsal surface of an entire hemisphere, the window size was approximately 6 x 3 mm2. Severe brain vibrations were not observed regardless of such a large window. Importantly, the condition of the brain surface did not deteriorate for more than one month. Wide-field imaging of a mouse expressing a genetically-encoded calcium indicator (GECI), GCaMP6f, specifically in astrocytes, revealed synchronized responses in a few millimeters. Two-photon imaging of the same mouse showed prominent calcium responses in individual astrocytes over several seconds. Furthermore, a thin layer of an adeno-associated virus was applied to the PVDC film and successfully expressed GECI in cortical neurons over the cranial window. This technique is reliable and cost-effective for making a large cranial window and facilitates the investigation of the neural and glial dynamics and their interactions during behavior at the macroscopic and microscopic levels.

In Vivo Wide-Field and Two-Photon Calcium Imaging from a Mouse Using a Large Cranial Window

The present protocol describes making a large (6 x 3 mm2) cranial window using food wrap, transparent silicone, and cover glass. This cranial window allows in vivo wide-field and two-photon calcium imaging experiments in the same mouse.

Wide-field calcium imaging from the mouse&#39;s neocortex allows one to observe cortex-wide neural activity related to various brain functions. On the ...

A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

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