All animal experiments were performed in accordance with guidelines approved by the IACUC at the University of Nebraska Medical Center.
1. Before surgery
<ol>
	<li>Prepare pipettes for viral injections. Pull borosilicate glass capillaries using a pipette puller and bevel the pipette at a 20&#176; angle. Sterilize the pipettes overnight.</li>
	<li>Prepare fresh pieces of sterile gel foam by cutting them into small squares. Submerge the gel foam in a sterile microfuge tube containing 0.5 mL of saline. Soak the gel foam in saline for at least 30 min before use.</li>
	<li>Prepare fresh pieces of sponge strips by cutting them into thin strips.</li>
	<li>Twenty minutes before surgery, inject 4-6-week-old GLAST-CreER/GCaMP6f mice intraperitoneally with 0.2 mg/kg dexamethasone to prevent brain swelling and 5 mg/kg carprofen to reduce inflammation. 
	&#8203;NOTE: To induce the recombination and expression of GCaMP6f in approximately 40% of the astrocytes, GLAST-CreER/GCaMP6f mice were injected with 100 mg/kg tamoxifen at 3 weeks for 5 consecutive days.</li>
</ol>
2. Start of surgery
<ol>
	<li>After 20 min, anesthetize the mice with 3% isoflurane with oxygen at a flow rate of 1 L/min for approximately 1.5 min.</li>
	<li>Once anesthetized, place the mouse on a stereotaxic frame that sits on a water re-circulating blanket to maintain a body temperature of 37 &#176;C throughout the surgery. Secure the mouse to the frame using the snout clamp and ear bars. Ensure that the snout clamp and ear bars are stable enough that the head will not move during the procedure, but not so tight that the skull is damaged.</li>
	<li>Maintain anesthesia with 1-1.5% isoflurane with oxygen at a flow rate of 1-2 L/min. Note that some animals may need more or less isoflurane to maintain sedation. Monitor the breathing of the animal as well as its reflexes to toe and tail pinches, and adjust the isoflurane appropriately.</li>
	<li>Apply eye ointment to each eye using a cotton tip applicator to prevent the eyes from drying out during the procedure.</li>
	<li>Using rodent trimmers, shave the hair from the neck to just past the eyes. Use caution to ensure that the whiskers of the animal are not accidentally trimmed.</li>
	<li>Clean the shaved area with one iodine prep pad, followed by one alcohol prep pad.</li>
	<li>Using sterile tissue forceps and surgical scissors, cut and remove the skin from the frontal sutures anterior to the bregma all the way posteriorly to lambda.</li>
	<li>Once the skin is removed, add approximately 0.1 mL of 1% xylocaine (lidocaine with epinephrine 1:200,000) to the skull to minimize bleeding of the skull.</li>
	<li>Using a sterile size 11 carbon steel surgical blade attached to a handle, gently scrape the connective tissue from the skull. Take extra care when removing the connective tissue near the edge of the skull, as any remaining tissue could prevent strong adhesion of the glass or head plate later on.</li>
	<li>Use sterile tissue forceps to remove loose connective tissue from the skull.</li>
	<li>Using sterile cotton tip applicators, clean off any remaining xylocaine from the skull.</li>
	<li>Once completed, thoroughly degrease the skull using a sterile cotton tip applicator dipped in acetone. Use compressed air to immediately blow-dry the skull.</li>
</ol>
3. Craniotomy
<ol>
	<li>Using a fine-point marker and a ruler, measure the appropriate size for the opening at the desired location. Confirm the size of the opening by comparing it to the size of the cover glass (3 or 5 mm in diameter); ensure that the opening is slightly smaller than the size of the cover glass.</li>
	<li>Using a dental drill, gradually trace the outline of the opening, thinning the bone. Drill slowly to prevent drilling through the bone, and drill in concentric circles to facilitate even thinning of the bone.</li>
	<li>After each concentric pass, add a drop or two of sterile saline to the drilled area. Allow the saline to sit for at least 10 s to prevent the bone from overheating and damaging the underlying dura.</li>
	<li>Use a sterile cotton tip applicator to absorb any remaining saline. Blow compressed air over the area to ensure that all remaining saline has evaporated before resuming drilling. Do this to blow away bone debris that remains.</li>
	<li>Repeat steps 3.2-3.4 until the bone is adequately thinned for removal.</li>
	<li>Ensure that the bone is adequately thinned for removal by gently pushing on the central area of the bone with fine forceps to check that the thinned skull moves. The underlying vasculature should be visible where the drilling occurred. Observe that the bone may appear as if it will crack where thinning occurred. 
	NOTE: Training at this stage is crucial to identify when the bone has been thinned appropriately.</li>
	<li>Before attempting to remove the bone, check the mouse to make sure it is completely sedated. If not, gradually increase the isoflurane maintenance to prevent brain swelling when removing the bone.</li>
	<li>Add a small piece of gel foam saturated in saline to the thinned skull. Add one or two additional drops of saline to the thinned skull. 
	NOTE: Having plenty of saline over the thinned skull helps to reduce bleeding and protect the dura when removing the bone flap.</li>
	<li>Using a miniature 15&#176;&#160;pointed blade, carefully insert the blade into the thinned bone and cut along the thinned bone. Keep the gel foam soaked in saline, ready to stop any bleeds that may occur.</li>
	<li>Using forceps, carefully lift and remove the bone. Be as gentle as possible to avoid damage to the underlying tissue and vasculature. Take extreme care not to damage the dura mater.</li>
	<li>Once the bone flap is removed, add a fresh piece of gel foam saturated in saline to the cortex. Add a few drops of saline to prevent the gel foam from drying out, as this will cause damage to the underlying tissue.</li>
</ol>
4. Viral injections
<ol>
	<li>Perform viral injections using a stereotaxic apparatus.
	<ol>
		<li>Prepare AAV1.CaMKII.0.4.Cre (titer of 3 &#215; 1013 genome copies (GC)/mL) at 1:5,000 by serial dilutions in saline.</li>
		<li>Prepare the mixture of AAV1.CaMKII.0.4.Cre (1:5,000) and AAV1.CAG.FLEX.tdTomato (titer of 5 &#215; 1012 GC/mL) on a small piece of parafilm.</li>
		<li>Fill a sterile beveled glass pipette with a tip size of 20 &#181;m with the virus mixture by gently placing the pipette on the solution.</li>
		<li>Lower the pipette so that it just touches the surface of the brain and continue to lower for an additional 200-300 &#181;m for L2/3 injections. Using an intracellular microinjection dispense system (see the Table of Materials), pressure-inject 12-15 times over 2 min (20 psi, 9 ms pulse duration). Observe the meniscus in the pipette drop with each injection to ensure that the pipette is not blocked.</li>
		<li>Once injected, leave the pipette in the brain for 4-5 min to prevent backflow. Slowly retract the pipette.</li>
		<li>Repeat the injections at 2-3 sites (approximately 500 &#181;m apart).</li>
		<li>Discard the used glass pipettes, parafilm, pipette tips, and microfuge tubes used for serial dilutions of the Cre virus into 10% bleach.</li>
	</ol>
	</li>
</ol>
5. Implantation of cranial window
<ol>
	<li>Remove any gel foam, and use a small strip of sponge strips to soak up any remaining saline.</li>
	<li>Add a few drops of the antibiotic enrofloxacin (22.7 &#181;g/mL) to the opening and allow it to remain there for 1 min. After 1 min, use a fresh piece of sponge strip to absorb any remaining enrofloxacin. Repeat this process two more times.</li>
	<li>After the third wash, add a few drops of saline to the opening and allow it&#160;to remain there for 1 min. After 1 min, use a fresh piece of sponge strip to absorb any remaining saline. Repeat this wash process two more times and after the third wash, add a small drop of saline to the opening.</li>
	<li>Place the cover glass over the opening. Use forceps to ensure that the glass is flush over the opening to obtain images of good quality.</li>
	<li>While gently holding the glass in place, apply cyanoacrylate adhesive gel (Table of Materials) around the perimeter of the glass to seal the edges of the window to the skull. Make sure not to let the glue seep under the cover glass, and keep as much glue off the surface of the cover glass as possible.</li>
	<li>Apply a layer of super glue over the adhesive gel. Add a layer of dental cement liquid over the glue to allow it to harden.</li>
	<li>Take an appropriately sized helicopter-type head plate and apply a thin layer of glue around the central opening. Place the head plate over the cover glass, and allow the glue to dry briefly.</li>
	<li>In a 1.5 mL microfuge tube, add dental cement powder to the 0.1 mL mark on the tube. Add 7-8 drops of fast-curing, instant adhesive, and mix. Draw into a 1 mL syringe with a 19 G needle that has been cut to create a larger opening.</li>
	<li>Inject the mixture through the lateral holes of the helicopter bar until it seeps from either side. Apply the dental cement/adhesive mixture to the rest of the exposed skull to fasten the headplate to the skull, which will reduce movement artifacts during imaging.</li>
	<li>Allow the mixture to air-dry for at least 15 min.</li>
	<li>Inject the mouse with 0.5 mL of saline subcutaneously to aid in recovery.</li>
</ol>
6. Post operation
<ol>
	<li>Remove the mouse from the stereotaxic frame and return it to its cage.</li>
	<li>Place a portion of the cage on a water re-circulating blanket, space gel heating pads or isothermal pads to assist with recovery.</li>
	<li>Provide the animal with a small helping of food pellets and wet food, in addition to their regular diet, to further aid in recovery. Add new food pellet and wet food each day after surgery for the duration of recovery.</li>
	<li>The day after surgery, inject mice once with 5 mg/kg carprofen and 5 mg/kg enrofloxacin.</li>
	<li>On days 2-6, inject the mice once with 5 mg/kg carprofen and twice with 5 mg/kg enrofloxacin, pending approval by the local IACUC. Separate the enrofloxacin injections by at least 8 h.</li>
	<li>On days 7-20, inject the mice daily with 5 mg/kg carprofen.</li>
</ol>
7. Animal habituation for imaging
<ol>
	<li>On day 14 after surgery, examine the cranial window for optical clarity. Do not use mice with unclear or otherwise damaged windows (i.e., excessive angiogenesis).</li>
	<li>Check for tdTomato expression under a fluorescence microscope. If labeled cells can be identified, proceed with animal habituation.</li>
	<li>First day of habituation (handling)
	<ol>
		<li>Hold the mouse for a few minutes and return it to its cage after handling. Repeat the handling three times with a 15 min interval between the sessions.</li>
		<li>After the third trial, habituate the mouse by wrapping the animal in a small piece of cloth.</li>
		<li>Hold the mouse wrapped in cloth for approximately 1 min.</li>
		<li>After the trial, return the mouse to its cage. Repeat this process two more times, allowing a 15 min interval between each trial.</li>
	</ol>
	</li>
	<li>Second day of habituation
	<ol>
		<li>Weigh and record the mouse weight.</li>
		<li>Wrap the mouse in cloth, and secure it via its head plate to the head fixation arm of the air-lifted home cage.</li>
		<li>Leave the home cage exposed to the light.</li>
		<li>Allow the mouse to remain secured in the mobile home cage for 15 min.</li>
		<li>After habituation, remove the mouse from the home cage and turn off the airflow.</li>
		<li>Weigh the mouse and return it to its cage. Take care to only include mice that do not lose more than 10% of their body weight. Exclude mice that lose more than 10% of their weight during any day of habituation from imaging experiments.</li>
	</ol>
	</li>
	<li>Day three of habituation and beyond
	<ol>
		<li>Weigh and record the mouse weight before securing it to the air-lifted home cage.</li>
		<li>Secure the mouse to the air-lifted home cage.</li>
		<li>Cover the home cage with something that will provide a dark interior, such as a box, and allow the mouse to remain secured for 30 min.</li>
		<li>After 30 min, uncover the home cage, remove the mouse, and turn off the air flow.</li>
		<li>Weigh and record the mouse weight after habituation.</li>
		<li>Repeat this process every day, increasing the duration the mouse is secured to the home cage by 15 min. Continue this process until the mouse can be secured to the home cage for 1.5 h as this is the approximate duration of a two-photon imaging session.</li>
		<li>To habituate the mouse to noise, perform some habituation sessions on the microscope to acclimate the mouse to the sounds of the laser scanning mirrors. Exclude mice that fail to habituate sufficiently (i.e., vocalizations and stress-induced defecation).</li>
	</ol>
	</li>
</ol>
8. Multiphoton imaging
<ol>
	<li>Commence imaging 3 weeks following surgery to allow for the window to clear.</li>
	<li>Perform imaging on a custom-made or commercially available multiphoton microscope equipped with a Ti:Sapphire laser. Acquire images using a high numerical aperture (NA) water immersion objective. 
	NOTE: Two-channel imaging is achieved by using a 565 nm dichroic mirror and two external photomultiplier tubes. A 535/50 bandpass filter is used to detect GCaMP6f emission, and a 610/75 bandpass filter is used to detect tdTomato. A 25x water immersion objective (1.05 NA) and resonant scanners were used to acquire images shown in Figure 1 and Figure 2. Each region of interest consisted of a stack of images separated axially by 1 &#181;m. Each optical section was collected at 512 x 512 pixels, 0.18&#160;&#181;m/pixel. Images were acquired from the forelimb region of the primary motor cortex as determined by stereotaxic coordinates.</li>
	<li>Set the wavelength of the laser to 920 nm (990 nm if a red-shifted GECI is used).</li>
	<li>Place the mouse on the mobile home cage, and clamp the headplate to the head fixation arm.</li>
	<li>Add a few drops of water to the center of the window.</li>
	<li>Using the wide-field mode of the microscope, select 2-3 positions of easily identifiable vasculature. Save the images of these blood vessels and record the X- and Y-coordinates that appear on the motor controller to relocate the tdTomato-labeled dendrites for subsequent imaging sessions. Adjust the X- and Y-coordinates each time to realign with the saved images of the blood vessels.</li>
	<li>Once the vasculature has been imaged and positions recorded, locate the regions with neurons expressing tdTomato (Figure 1). Image the synaptic structures (dendrites and axons) of neurons and GCaMP6f activity in astrocytes. 
	NOTE: The dendrites will serve as landmarks to reliably return to the same regions and image the same dendrites and astrocytes over repeated days. No detectable shift in the imaging plane was observed with stable head fixation and after habituation to head fixation. Displacement that occurs in the X and Y directions is motion-corrected.</li>
	<li>After imaging, return the mouse to its cage. 
	NOTE: Mice are typically kept on the scope for a maximum of 2 h. When the imaging lasts for a long time, the water under the objective may dry. Water should be added during the experiments. Alternatively, an ultrasonic gel can be used instead of water.</li>
</ol>

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(88), e51869 (2014).</li><li>Holtmaat, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=high-resolution+imaging+in+the+mouse+neocortex+through+a+chronic+cranial+window.">high-resolution imaging in the mouse neocortex through a chronic cranial window.</a> Nature Protocols. 4 (8), 1128-1144 (2009).</li><li>Hauglund, N. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Meningeal+lymphangiogenesis+and+enhanced+glymphatic+activity+in+mice+with+chronically+implanted+EEG+electrodes.">Meningeal lymphangiogenesis and enhanced glymphatic activity in mice with chronically implanted EEG electrodes.</a> Journal of Neuroscience. 40 (11), 2371-2380 (2020).</li><li>De Paola, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+type-specific+structural+plasticity+of+axonal+branches+and+boutons+in+the+adult+neocortex.">Cell type-specific structural plasticity of axonal branches and boutons in the adult neocortex.</a> Neuron. 49 (6), 861-875 (2006).</li><li>Cheng, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+two-photon+calcium+imaging+at+different+depths+with+spatiotemporal+multiplexing.">Simultaneous two-photon calcium imaging at different depths with spatiotemporal multiplexing.</a> Nature Methods. 8 (2), 139-142 (2011).</li><li>Yang, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thinned-skull+cranial+window+technique+for+long-term+imaging+of+the+cortex+in+live+mice.">Thinned-skull cranial window technique for long-term imaging of the cortex in live mice.</a> Nature Protocols. 5 (2), 201-208 (2010).</li><li>Shih, A. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+polished+and+reinforced+thinned-skull+window+for+long-term+imaging+of+the+mouse+brain.">A polished and reinforced thinned-skull window for long-term imaging of the mouse brain.</a> Journal of Visualized Experiments: JoVE. (61), e3742 (2012).</li><li>Helm, P. J., Ottersen, O. P., Nase, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+optical+properties+of+the+mouse+cranium--implications+for+in+vivo+multi+photon+laser+scanning+microscopy.">Analysis of optical properties of the mouse cranium--implications for in vivo multi photon laser scanning microscopy.</a> Journal of Neuroscience Methods. 178 (2), 316-322 (2009).</li><li>Stobart, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+in+vivo+calcium+imaging+of+astrocytes+reveals+distinct+cellular+compartment+responses+to+sensory+stimulation.">Long-term in vivo calcium imaging of astrocytes reveals distinct cellular compartment responses to sensory stimulation.</a> Cerebral Cortex. 28 (1), 184-198 (2018).</li><li>Pryazhnikov, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Longitudinal+two-photon+imaging+in+somatosensory+cortex+of+behaving+mice+reveals+dendritic+spine+formation+enhancement+by+subchronic+administration+of+low-dose+ketamine.">Longitudinal two-photon imaging in somatosensory cortex of behaving mice reveals dendritic spine formation enhancement by subchronic administration of low-dose ketamine.</a> Scientific Reports. 8 (1), 6464 (2018).</li><li>Thrane, A. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=General+anesthesia+selectively+disrupts+astrocyte+calcium+signaling+in+the+awake+mouse+cortex.">General anesthesia selectively disrupts astrocyte calcium signaling in the awake mouse cortex.</a> Proceedings of the National Academy of Sciences of the United States of America. 109 (46), 18974-18979 (2012).</li><li>Paukert, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Norepinephrine+controls+astroglial+responsiveness+to+local+circuit+activity.">Norepinephrine controls astroglial responsiveness to local circuit activity.</a> Neuron. 82 (6), 1263-1270 (2014).</li><li>Delekate, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metabotropic+P2Y1+receptor+signalling+mediates+astrocytic+hyperactivity+in+vivo+in+an+Alzheimer's+disease+mouse+model.">Metabotropic P2Y1 receptor signalling mediates astrocytic hyperactivity in vivo in an Alzheimer's disease mouse model.</a> Nature Communications. 5, 5422 (2014).</li></ol>

Here, we have presented a protocol for the implantation of chronic cranial windows for in vivo imaging of cortical astrocytes and neurons in awake, head-restrained mice on an air-lifted home cage. Specific examples have been provided of the cranial window application for imaging astrocytes that express GECIs and neuronal synaptic structures. With the use of multiphoton microscopy, astrocytic calcium signaling dynamics and structural synaptic dynamics can be recorded repeatedly over days.
<p class="jove_conte...

The ability to perform in vivo multiphoton fluorescence microscopy in the cortex of mice is paramount to the study of cellular signaling and structure1,2,3,4,5,6,7,8,9, disease pathology10,11, and cellular development12,13. With the implantation of chronic cranial windows, longitudinal imaging is possible, allowing for repeated imaging of cortical areas for days, weeks, or months13,14 in live animals. Multiphoton microscopy is ideal for in vivo, repeated imaging because of improved depth probing and reduced photodamage associated with the infrared laser used. This allows for the study of molecular and cellular dynamics of specific cells in various cortical regions.
Multiphoton microscopy has been used for in vivo imaging of neuronal and glial cells in mice15,16,17,18,19,20. Various strategies can be implemented to label particular cell types and areas of interest. One common approach is to drive the expression of genetically encoded fluorescent proteins in a cell-specific manner using the Cre-Lox recombination system. This can be performed with genetically modified mice, e.g., crossing tdTomato &#34;floxed&#34; mouse (Ai14) with a mouse expressing Cre-recombinase under a promoter of interest21. Alternatively, cell- and site-specific labeling can be achieved with viral injections. Here, a virus encoding Cre recombinase under a cell-specific promotor and a virus encoding a floxed gene of interest are injected into a defined region. Appropriate cell types receiving both viral vectors will then express the desired gene(s). These genes can be structural markers, such as tdTomato, to view changes in cellular morphology22 or genetically encoded calcium indicators (GECIs), such as GCaMP and/or RCaMP, to examine calcium dynamics23. Methods of genetic recombination can be applied individually or in combination to label one or more cell types. A third approach, not requiring transgenic mice or viral constructs (which have limited packaging capacity), is in utero electroporation of DNA constructs24. Depending on the timing of the electroporation, different cell types can be targeted25,26,27.
When performing multiphoton imaging, mice can be imaged while awake or anesthetized. Imaging of awake mice can be performed by securing the mouse via an attached head plate28. This approach is made less stressful by allowing relatively free movement of the animal using methods, such as free-floating, air-supported Styrofoam balls29, free-floating treadmills1, or an air-lifted home cage system where mice are fastened by an attached head plate and allowed to move in an open chamber30. For each of these imaging conditions, it will first be necessary to habituate the mice to the imaging setup. This paper describes the habituation and imaging procedure using an air-lifted home cage system.
This protocol describes the implantation of a chronic cranial window for longitudinal in vivo imaging in the cortex. Here, we will use mice that conditionally express GCaMP6f in astrocytes to monitor calcium signaling dynamics. Further, this paper describes the procedure for viral injections using tdTomato as a label for neurons. This allows the determination of changes in neuronal synaptic structure and/or the availability as a structural marker that enables repeated imaging of the same astrocyte. Throughout the protocol, crucial steps will be highlighted to ensure the best possible quality of images obtained from multiphoton microscopy.

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			<td>A16P4</td>
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			<td>Alcohol Prep Pads</td>
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			<td>Nikon</td>
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			<td>Patterson Dental</td>
			<td>602-7932</td>
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			<td>Bayer</td>
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			<td>Dechra</td>
			<td>17033-211-38</td>
			<td>Surgery Supply</td>
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			<td>Dolan-Jenner Industries</td>
			<td></td>
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			<td>World Precision Instruments</td>
			<td>14099</td>
			<td>Surgery Tools</td>
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			<td>World Precision Instruments</td>
			<td>501764</td>
			<td>Surgery Tools</td>
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		<tr>
			<td>GCaMP6f B6; 129S-Gt(ROSA)26Sortm95.1(CAGGCaMP6f)Hze/J</td>
			<td>The Jackson Laboratory</td>
			<td>Stock No: 024105</td>
			<td>Mouse line</td>
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		<tr>
			<td>Germinator</td>
			<td>Fisher Scientific</td>
			<td></td>
			<td>Equipment</td>
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			<td>GLAST-CreER Tg(Slc1a3-cre/ERT) 1Nat/J</td>
			<td>The Jackson Laboratory</td>
			<td>Stock No: 012586</td>
			<td>Mouse Line</td>
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			<td>Headplate</td>
			<td>Neurotar</td>
			<td>Model 1, Model 3</td>
			<td>Surgery Supply</td>
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			<td>Hemostatic forceps</td>
			<td>World Precision Instruments</td>
			<td>501705</td>
			<td>Surgery Tools</td>
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			<td>World Precision Instruments</td>
			<td>501247</td>
			<td>Surgery Tools</td>
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			<td>World Precision Instruments</td>
			<td>500236</td>
			<td>Surgery Tools</td>
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			<td>Avantor</td>
			<td>15648-926</td>
			<td>Surgery Supply</td>
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			<td>Piramal</td>
			<td></td>
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			<td>Isoflurane table top system with Induction Box</td>
			<td>Harvard Apparatus</td>
			<td></td>
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			<td>SurgiVet</td>
			<td></td>
			<td>Equipment</td>
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			<td>Henkel</td>
			<td>40140</td>
			<td>fast-curing instant adhesive</td>
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			<td>NC9194415</td>
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			<td>Sutter Instruments</td>
			<td></td>
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			<td>Multiphoton Microscope</td>
			<td></td>
			<td></td>
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			<td>Matheson</td>
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			<td>Matheson</td>
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			<td>Parker</td>
			<td></td>
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			<td>17014340</td>
			<td>Surgery Supply</td>
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			<td>Fisher Scientific</td>
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		</tr>
	</tbody>
</table>

implantation of a cranial window for repeated in vivo imaging in awake mice

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their activity in vivo in behaving mice. This paper describes a method for the implantation of a chronic cranial window to allow for the longitudinal imaging of brain cells in awake, head-restrained mice. In combination with genetic strategies and viral injections, it is possible to label specific cells and regions of interest with structural or physiological markers. This protocol demonstrates how to combine viral injections to label neurons in the vicinity of GCaMP6-expressing astrocytes in the cortex for simultaneous imaging of both cells through a cranial window. Multiphoton imaging of the same cells can be performed for days, weeks, or months in awake, behaving animals. This approach provides researchers with a method for viewing cellular dynamics in real time and can be applied to answer a number of questions in neuroscience.

The quality of the cranial window can be assessed by how crisp the neuronal structures appear. In a good window, dendritic spines are clearly visible (Figure 1). With the structural and positional data stored, the same animal can be imaged repeatedly for days, weeks, or months to examine the same cells (Figure 1). The images in Figure 1 were obtained from the forelimb region of the primary motor cortex (in a 5 mm window). A variety ...

Watch this Scientific Journal Video about Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice at JoVE.com

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

Presented here is a protocol for the implantation of a chronic cranial window for the longitudinal imaging of brain cells in awake, head-restrained mice.

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their ...

implantation-cranial-window-for-repeated-vivo-imaging-awake

Research

JoVE Journal

Neuroscience

7.0K Views.  University of Nebraska Medical Center. Presented here is a protocol for the implantation of a chronic cranial window for the longitudinal imaging of brain cells in awake, head-restrained mice.

Video: Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or thinned-skull 3 preparations. While the open-skull technique is very versatile, it is not optimal for studying microglia because it is invasive and can cause microglial activation. Even though the thinned-skull approach is minimally invasive, the repeated re-thinning of skull required for chronic imaging increases the risks of tissue injury and microglial activation and allows for a limited number of imaging sessions. Here we present a chronic thin-skull window method for monitoring murine microglia in vivo over an extended period of time using two-photon microscopy. We demonstrate how to prepare a stable, accessible, thinned-skull cortical window (TSCW) with an apposed glass coverslip that remains translucent over the course of three weeks of intermittent observation. This TSCW preparation is far more immunologically inert with respect to microglial activation than open craniotomy or repeated skull thinning and allows an arbitrary number of imaging sessions during a time period of weeks. We prepare TSCW in CX3CR1 GFP/+ mice 4 to visualize microglia with enhanced green fluorescent protein to &le;150 &mu;m beneath the pial surface. We also show that this preparation can be used in conjunction with stereotactic brain injections of the HIV-1 neurotoxic protein Tat, adjacent to the TSCW, which is capable of inducing durable microgliosis. Therefore, this method is extremely useful for examining changes in microglial morphology and motility over time in the living brain in models of HIV Associated Neurocognitive Disorder (HAND) and other neurodegenerative diseases with a neuroinflammatory component.

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

We describe a method for repeatedly visualizing murine microglia and circulating monocytes in vivo over hours, days or weeks using transcranial two-photon microscopy. We demonstrate how to prepare a thinned-skull window that allows intermittent observation of quiescent microglia that can be activated by adjacent stereotactic injection of the HIV-1 regulatory protein Tat.

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

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 ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

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 ...

Piezo High Accuracy Surgical Osteal Removal (PHASOR): A Technique for Improved Cranial Window Surgery in Mice

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated thinning of the skull. Cranial window surgery is typically performed with a high speed rotary drill, and many investigators find it challenging to prevent the drill from damaging the delicate dura and blood vessels. Extensive training and practice is required to remove the bone without damage to underlying tissue and thus cranial window surgery can be difficult, time consuming, and produce tissue damage. Piezoelectric surgery, which is extensively used for maxillofacial and dental surgery, utilizes ultrasonic vibrations to remove bone without damaging soft tissues. We have developed a method applying piezoelectric surgery to improve cranial window surgery in mice in preparation for multiphoton imaging. Comparisons within our lab find that the method requires less surgery time and has a lower average rate of complications due to dural bleeding than cranial window surgery with a rotary drill.

Piezo High Accuracy Surgical Osteal Removal PHASOR: A Technique for Improved Cranial Window Surgery in Mice

Piezoelectric surgery has led to improvements in human maxillofacial and dental surgery. We have developed a protocol to optimize piezoelectric surgery for cranial window surgery in mice.

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the ...

Construction and Implementation of Carbon Fiber Microelectrode Arrays for Chronic and Acute In Vivo Recordings

Multichannel electrode arrays offer insight into the working brain and serve to elucidate neural processes at the single-cell and circuit levels. Development of these tools is crucial for understanding complex behaviors and cognition and for advancing clinical applications. However, it remains a challenge to densely record from cell populations stably and continuously over long time periods. Many popular electrodes, such as tetrodes and silicon arrays, feature large cross-diameters that produce damage upon insertion and elicit chronic reactive tissue responses associated with neuronal death, hindering the recording of stable, continuous neural activity. In addition, most wire bundles exhibit broad spacing between channels, precluding simultaneous recording from a large number of cells clustered in a small area. The carbon fiber microelectrode arrays described in this protocol offer an accessible solution to these concerns. The study provides a detailed method for fabricating carbon fiber microelectrode arrays that can be used for both acute and chronic recordings in vivo. The physical properties of these electrodes make them ideal for stable and continuous long-term recordings at high cell densities, enabling the researcher to make robust, unambiguous recordings from single units across months.

Construction and Implementation of Carbon Fiber Microelectrode Arrays for Chronic and Acute In Vivo Recordings

This protocol describes a procedure for constructing carbon fiber microelectrode arrays for chronic and acute in vivo electrophysiological recordings in mouse (Mus musculus) and ferret (Mustela putorius furo) from multiple brain regions. Each step, following the purchase of raw carbon fibers to microelectrode array implantation, is described in detail, with emphasis on microelectrode array construction.

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

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A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

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

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

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

Piezo High Accuracy Surgical Osteal Removal (PHASOR): A Technique for Improved Cranial Window Surgery in Mice

Laminectomy and Spinal Cord Window Implantation in the Mouse

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

In vivo Calcium Imaging in Mouse Inferior Olive

Construction and Implementation of Carbon Fiber Microelectrode Arrays for Chronic and Acute In Vivo Recordings