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><tbody><tr><td>Heating Pad</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&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;mg/mL</td><td>Western Drug Distribution Centre Ltd</td><td>125231</td><td></td></tr><tr><td>Butyl cyanoacrylate glue (VetBond)</td><td>Western Drug Distribution Centre Ltd</td><td>12612</td><td></td></tr></tbody></table>

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, ...

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Research

JoVE Journal

Neuroscience

12.2K Views.  University of Lethbridge. 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.

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

A Craniotomy Surgery Procedure for Chronic Brain Imaging

Imaging techniques are becoming increasingly important in the study brain function. Among them, two-photon laser scanning microscopy has emerged as an extremely useful method, because it allows the study of the live intact brain. With appropriate preparations, this technique allows the observation of the same cortical area chronically, from minutes to months. In this video, we show a preparation for chronic in vivo imaging of the brain using two-photon microscopy. This technique was initially pioneered by Dr. Karel Svoboda, who is now a Howard Hughes Medical Institute Investigator at Janelia Farm. Preparations like the one shown here can be used for imaging of neocortical structure (e.g., dendritic and axonal dynamics),  to record neuronal activity using calcium-sensitive dyes, to image cortical blood flow dynamics, or for intrinsic optical imaging studies. Deep imaging of the neocortex is possible with optimal cranial window surgeries. Operating under the most sterile conditions possible to avoid infections, together with using extreme care to do not damage the dura mater during the surgery, will result in successful and long-lasting glass-covered cranial windows.


This video and protocol demonstrate how to implant a glass-covered cranial window in rodents. These preparations can be used for chronic in vivo two-photon imaging of the neocortex over time scales of months. It may also be used for other types of imaging, including optical intrinsic signal imaging.

Imaging techniques are becoming increasingly important in the study brain function. Among them, two-photon laser scanning microscopy has emerged as an ...

Biology

Longitudinal Intravital Imaging of Brain Tumor Cell Behavior in Response to an Invasive Surgical Biopsy

Biopsies are standard of care for cancer treatment and are clinically beneficial as they allow solid tumor diagnosis, prognosis, and personalized treatment determination. However, perturbation of the tumor architecture by biopsy and other invasive procedures has been associated with undesired effects on tumor progression, which need to be studied in depth to further improve the clinical benefit of these procedures. Conventional static approaches, which only provide a snapshot of the tumor, are limited in their ability to reveal the impact of biopsy on tumor cell behavior such as migration, a process closely related to tumor malignancy. In particular, tumor cell migration is the key in highly aggressive brain tumors, where local tumor dissemination makes total tumor resection virtually impossible. The development of multiphoton imaging and chronic imaging windows allows scientists to study this dynamic process in living animals over time. Here, we describe a method for the high-resolution longitudinal imaging of brain tumor cells before and after a biopsy in the same living animal. This approach makes it possible to study the impact of this procedure on tumor cell behavior (migration, invasion, and proliferation). Furthermore, we discuss the advantages and limitations of this technique, as well as the ability of this methodology to study changes in the cancer cell behavior for other surgical interventions, including tumor resection or the implantation of chemotherapy wafers.

Here we describe a method for high-resolution time-lapse multiphoton imaging of brain tumor cells before and after invasive surgical intervention (e.g., biopsy) within the same living animal. This method allows studying the impact of these invasive surgical procedures on tumor cells&#39; migratory, invasive, and proliferative behavior at a single cell level.

Biopsies are standard of care for cancer treatment and are clinically beneficial as they allow solid tumor diagnosis, prognosis, and personalized ...

Cancer Research

Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, direction5-7, ocular dominance8,9 and binocular disparity9. Orientation selectivity is the most widely studied feature and a continuous map with a quasi-periodic layout for preferred orientation is present across the entire primary visual cortex10,11. Integrating the synaptic, cellular and network contributions that lead to stimulus selective responses in these functional maps requires the hybridization of imaging techniques that span sub-micron to millimeter spatial scales. With conventional intrinsic signal optical imaging, the overall layout of functional maps across the entire surface of the visual cortex can be determined12. The development of in vivo two-photon microscopy using calcium sensitive dyes enables one to determine the synaptic input arriving at individual dendritic spines13 or record activity simultaneously from hundreds of individual neuronal cell bodies6,14. Consequently, combining intrinsic signal imaging with the sub-micron spatial resolution of two-photon microscopy offers the possibility of determining exactly which dendritic segments and cells contribute to the micro-domain of any functional map in the neocortex. Here we demonstrate a high-yield method for rapidly obtaining a cortical orientation map and targeting a specific micro-domain in this functional map for labeling neurons with fluorescent dyes in a non-rodent mammal. With the same microscope used for two-photon imaging, we first generate an orientation map using intrinsic signal optical imaging. Then we show how to target a micro-domain of interest using a micropipette loaded with dye to either label a population of neuronal cell bodies or label a single neuron such that dendrites, spines and axons are visible in vivo. Our refinements over previous methods facilitate an examination of neuronal structure-function relationships with sub-cellular resolution in the framework of neocortical functional architectures.

A method is described for labeling neurons with fluorescent dyes in predetermined functional micro-domains of the neocortex. First, intrinsic signal optical imaging is used to obtain a functional map. Then two-photon microscopy is used to label and image neurons within a micro-domain of the map.

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, ...

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

Optical coherence tomography (OCT) is a biomedical imaging technique with high spatial-temporal resolution. With its minimally invasive approach OCT has been used extensively in ophthalmology, dermatology, and gastroenterology1-3. Using a thinned-skull cortical window (TSCW), we employ spectral-domain OCT (SD-OCT) modality as a tool to image the cortex in vivo. Commonly, an opened-skull has been used for neuro-imaging as it provides more versatility, however, a TSCW approach is less invasive and is an effective mean for long term imaging in neuropathology studies. Here, we present a method of creating a TSCW in a mouse model for in vivo OCT imaging of the cerebral cortex.

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

We present a method of creating a thinned-skull cortical window (TSCW) in a mouse model for in vivo OCT imaging of the cerebral cortex.

Optical coherence tomography (OCT) is a biomedical imaging technique  with high spatial-temporal resolution. With its minimally invasive approach OCT  has ...

Laser Nanosurgery of Cerebellar Axons In Vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to understand the mechanism that promotes neuronal regeneration, an in-depth analysis of the neuronal types that can remodel after injury is needed. Several studies showed that damaged climbing fibers are capable of regrowing also in adult animals1,2. The investigation of the time-lapse dynamics of degeneration and regeneration of these axons within their complex environment can be performed by time-lapse two-photon fluorescence (TPF) imaging in vivo3,4. This technique is here combined with laser surgery, which proved to be a highly selective tool to disrupt fluorescent structures in the intact mouse cortex5-9.
This protocol describes how to perform TPF time-lapse imaging and laser nanosurgery of single axonal branches in the cerebellum in vivo. Olivocerebellar neurons are labeled by anterograde tracing with a dextran-conjugated dye and then monitored by TPF imaging through a cranial window. The terminal portion of their axons are then dissected by irradiation with a Ti:Sapphire laser at high power. The degeneration and potential regrowth of the damaged neuron are monitored by TPF in vivo imaging during the days following the injury.

Laser Nanosurgery of Cerebellar Axons In Vivo

Two-photon imaging, coupled to laser nanodissection, are useful tools to study degenerative and regenerative processes in the central nervous system with subcellular resolution. This protocol shows how to label, image, and dissect single climbing fibers in the cerebellar cortex in vivo.

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to ...

Lateral Chronic Cranial Window Preparation Enables In Vivo Observation Following Distal Middle Cerebral Artery Occlusion in Mice

Focal cerebral ischemia (i.e., ischemic stroke) may cause major brain injury, leading to a severe loss of neuronal function and consequently to a host of motor and cognitive disabilities. Its high prevalence poses a serious health burden, as stroke is among the principal causes of long-term disability and death worldwide1. Recovery of neuronal function is, in most cases, only partial. So far, treatment options are very limited, in particular due to the narrow time window for thrombolysis2,3. Determining methods to accelerate recovery from stroke remains a prime medical goal; however, this has been hampered by insufficient mechanistic insights into the recovery process. Experimental stroke researchers frequently employ rodent models of focal cerebral ischemia. Beyond the acute phase, stroke research is increasingly focused on the sub-acute and chronic phase following cerebral ischemia. Most stroke researchers apply permanent or transient occlusion of the MCA in mice or rats. In patients, occlusions of the MCA are among the most frequent causes of ischemic stroke4. Besides proximal occlusion of the MCA using the filament model, surgical occlusion of the distal MCA is probably the most frequently used model in experimental stroke research5. Occlusion of a distal (to the branching of the lenticulo-striate arteries) MCA branch typically spares the striatum and primarily affects the neocortex. Vessel occlusion can be permanent or transient. High reproducibility of lesion volume and very low mortality rates with respect to the long-term outcome are the main advantages of this model. Here, we demonstrate how to perform a chronic cranial window (CW) preparation lateral to the sagittal sinus, and afterwards how to surgically induce a distal stroke underneath the window using a craniotomy approach. This approach can be applied for sequential imaging of acute and chronic changes following ischemia via epi-illuminating, confocal laser scanning, and two-photon intravital microscopy.

Lateral Chronic Cranial Window Preparation Enables In Vivo Observation Following Distal Middle Cerebral Artery Occlusion in Mice

Surgical occlusion of a distal middle cerebral artery branch (MCAo) is a frequently used model in experimental stroke research. This manuscript describes the basic technique of permanent MCAo, combined with the insertion of a lateral cranial window, which offers the opportunity for longitudinal intravital microscopy in mice.

Focal cerebral ischemia (i.e., ischemic stroke) may cause major brain injury, leading to a severe loss of neuronal function and consequently to a host of ...

Longitudinal In Vivo Imaging of the Cerebrovasculature: Relevance to CNS Diseases

Remodeling of the brain vasculature is a common trait of brain pathologies. In vivo imaging techniques are fundamental to detect cerebrovascular plasticity or damage occurring overtime and in relation to neuronal activity or blood flow. In vivo two-photon microscopy allows the study of the structural and functional plasticity of large cellular units in the living brain. In particular, the thinned-skull window preparation allows the visualization of cortical regions of interest (ROI) without inducing significant brain inflammation. Repetitive imaging sessions of cortical ROI are feasible, providing the characterization of disease hallmarks over time during the progression of numerous CNS diseases. This technique accessing the pial structures within 250 &#956;m of the brain relies on the detection of fluorescent probes encoded by genetic cellular markers and/or vital dyes. The latter (e.g., fluorescent dextrans) are used to map the luminal compartment of cerebrovascular structures. Germane to the protocol described herein is the use of an in vivo marker of amyloid deposits, Methoxy-O4, to assess Alzheimer&#39;s disease (AD) progression. We also describe the post-acquisition image processing used to track vascular changes and amyloid depositions. While focusing presently on a model of AD, the described protocol is relevant to other CNS disorders where pathological cerebrovascular changes occur.

Longitudinal In Vivo Imaging of the Cerebrovasculature: Relevance to CNS Diseases

This manuscript describes a procedure to track the remodeling of the cerebrovasculature during amyloid plaque accumulation in vivo using longitudinal two-photon microscopy. A thinned-skull preparation enables the visualization of fluorescent dyes to assess the progression of cerebrovascular damage in a mouse model of Alzheimer&#39;s disease.

Remodeling of the brain vasculature is a common trait of brain pathologies. In vivo imaging techniques are fundamental to detect cerebrovascular ...

Medicine

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 Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy (pCLE) of Hippocampal Capillaries in Awake Mice

The goal of this protocol is to describe fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE) in its specific application to elucidate capillary blood flow effects during seizures, driven by mural cells. In vitro and in vivo cortical imaging have shown that capillary constrictions driven by pericytes can result from functional local neural activity, as well as from drug application, in healthy animals. Here, a protocol is presented on how to use pCLE to determine the role of microvascular dynamics in neural degeneration in epilepsy, at any tissue depth (specifically in the hippocampus). We describe a head restraint technique that has been adapted to record pCLE in awake animals, to address potential side-effects of anesthetics on neural activity. Using these methods, electrophysiological and imaging recordings can be conducted over several hours in deep neural structures of the brain.

In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy pCLE of Hippocampal Capillaries in Awake Mice

Whereas multiphoton imaging is only effective at limited depths from the tissue&#8217;s surface, it is possible to achieve 3 &#181;m resolution imaging at any depth via pCLE. Here, we present a protocol to conduct pCLE imaging to measure microvascular dynamics in the hippocampus of ictal and wild-type mice.

The goal of this protocol is to describe fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE) in its specific application ...

Multilayer Imaging with Microprism in Head-Fixed and Freely Moving Animals

Long-Term Imaging of Identified Neural Populations using Microprisms in Freely Moving and Head-Fixed Animals

With the advancement of multi-photon microscopy and molecular technologies, fluorescence imaging is rapidly growing to become a powerful approach for studying the structure, function, and plasticity of living brain tissues. In comparison to conventional electrophysiology, fluorescence microscopy can capture the neural activity as well as the morphology of the cells, enabling long-term recordings of the identified neuron populations at single-cell or subcellular resolution. However, high-resolution imaging typically requires a stable, head-fixed setup that restricts the movement of the animal, and the preparation of a flat surface of transparent glass allows visualization of neurons at one or more horizontal planes but is limited in studying the vertical processes running across different depths. Here, we describe a procedure to combine a head plate fixation and a microprism that gives multilayer and multimodal imaging. This surgical preparation not only gives access to the entire column of the mouse visual cortex but allows two-photon imaging in a head-fixed position and one-photon imaging in a freely moving paradigm. Using this approach, one can sample identified cell populations across different cortical layers, register their responses under head-fixed and freely moving states, and track the long-term changes over months. Thus, this method provides a comprehensive assay of the microcircuits, enabling direct comparison of neural activities evoked by well-controlled stimuli and under a natural behavioral paradigm.

Author Spotlight: Comparative Imaging of Neural Activity in Awake and Freely Moving States

When integrated with a head-plate and an optical design compatible with both single- and two-photon microscopes, the microprism lens presents a significant advantage in measuring neural responses in a vertical column under diverse conditions, including well-controlled experiments in head-fixed states or natural behavioral tasks in freely moving animals.

With the advancement of multi-photon microscopy and molecular technologies, fluorescence imaging is rapidly growing to become a powerful approach for ...

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

Summary

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A Craniotomy Surgery Procedure for Chronic Brain Imaging

Longitudinal Intravital Imaging of Brain Tumor Cell Behavior in Response to an Invasive Surgical Biopsy

Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

Laser Nanosurgery of Cerebellar Axons In Vivo

Lateral Chronic Cranial Window Preparation Enables In Vivo Observation Following Distal Middle Cerebral Artery Occlusion in Mice

Longitudinal In Vivo Imaging of the Cerebrovasculature: Relevance to CNS Diseases

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy (pCLE) of Hippocampal Capillaries in Awake Mice

Long-Term Imaging of Identified Neural Populations using Microprisms in Freely Moving and Head-Fixed Animals