S.E. Lutz is supported by the National Center for Advancing Translational Sciences, National Institutes of Health, under Grant KL2TR002002 and University of Illinois Chicago College of Medicine start-up funds. Simon Alford is supported by RO1 MH084874. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH. The authors thank Dritan Agalliu in the Department of Neurology at Columbia University Medical Center for the Tg eGFP:Claudin-5 mice, scientific discussions, and insights into the development of the surgical protocol and imaging applications. The authors thank Sunil P. Gandhi in the Department of Neurobiology and Behavior at University of California, Irvine for designing the first prototype of the stereotactic apparatus and animal temperature controller, discussion of the surgical protocol, and training in two-photon microscopy. The authors also thank Steve Pickens (W. Nuhsbaum, Inc.) for assistance in customizing the surgical stereomicroscope, and Ron Lipinski (Whale Manufacturing) for machining stereotactic parts.

All experiments follow the University of Illinois, Chicago Institutional Animal Care and Use Committee protocols. This is a terminal procedure.&#160;
1. Reagent Preparation
<ol>
	<li>Prepare artificial cerebral spinal fluid (aCSF) to contain 125 mM NaCl, 5 mM KCl, 10 mM Glucose, 10 mM HEPES, 2 mM MgCl2&#183;6H2O, 2 mM CaCl2&#183;2H2O in ddH2O. Sterile filter and freeze in individual-use aliquots. Warm aCSF in a water bath to 39 &#176;C before use.</li>
	<li>Warm low melting-point agarose (2%) in aCSF until fully dissolved in a water bath set to 65 &#176;C. During the laminectomy, cool the melted agarose aliquot to 39 &#176;C in a water bath, so that it can ready at close to physiologic temperature for step 5.2. 
	Note: The agarose solution can be stored at -20 &#176;C in single-use aliquots.</li>
	<li>Prepare sterile 50 mg/mL carprofen in bacteriostatic water. Store at 4 &#176;C.</li>
	<li>Clean cover-glasses with 70% ethanol, three washes of ddH2O, and store dry in a dust-free container.</li>
</ol>
2. Backplate 3D Printing
<ol>
	<li>Use 3D CAD software is used to create a model to the dimensions shown in Figure 1. The interior is an ellipse widest at the bottom surface with respect to the printer and cut with a lofted cut to a smaller ellipse forming a lumen at the opposite surface. Two projecting arms with holes to accept screws extend laterally, for attachment to the backplate holding fork. From this 3D structure, create a triangulated 3D mesh file (.STL file). 
	Note: See Figure 1B&#8210;D and Supplementary Files 1 and 2.</li>
	<li>Upload the triangulated 3D mesh file to a 3D printer.</li>
	<li>Print backplates using a 0.4 mm hot-end nozzle and a 0.2 mm layer height. Select nozzle temperature of 205 &#176;C, bed temperature of 45 &#176;C, and printing speed of 45 mm/s.</li>
	<li>Assess the resultant 3D printed backplates visually for structural integrity (Figure 1E); gross structural failure (absent lumen, collapsed wall) indicates printing defects (Figure 1F).</li>
</ol>
3. Surgical Preparation
<ol>
	<li>Preheat the heating pad.</li>
	<li>Load isoflurane into the delivery syringe while working in a chemical fume hood. Attach the delivery syringe to the isoflurane unit.</li>
	<li>Select an 8&#8210;12-week-old mouse. Weigh the animal. Induce anesthesia using 2% isoflurane in an induction chamber. Inject carprofen at 5 mg/kg subcutaneously.</li>
	<li>Position the nose-cone and deliver isoflurane at 2% with a flow rate of 150 mL/min for maintenance of a surgical plane of anesthesia (Figure 2A&#8210;E). Wrap the heating pad with a disposable absorbent pad for ease of cleanup.</li>
	<li>Position an animal on the heating pad at the surgical station and install the nose cone. Lubricate the thermometer with petroleum jelly and insert it 5 mm into the rectum. Tape the thermometer probe to the tail for stability. Apply ophthalmic ointment to eyes of the mouse.</li>
	<li>To maintain hydration, apply 200 &#181;L of lactated Ringer&#8217;s solution by subcutaneous injection every 30 min until termination of the experiment.</li>
	<li>Spray the dorsum with 70% ethanol, remove fur with clippers, and clean the site with povidone-iodine.</li>
</ol>
4. Laminectomy
<ol>
	<li>Position the animal between the ear bars; these maintain the head position of the mouse with respect to the nose-cone.</li>
	<li>Confirm that animal is deeply anesthetized as assessed by lack of interdigital pinch reflex and steady respiratory pattern.</li>
	<li>Make a 1.5 cm rostral-caudal incision at midline over the lower thoracic/upper lumbar region using #11 blade (Figure 2). Separate the skin by grasping it with blunt toothed forceps and/or gloved fingers. Use forceps to separate and peel back any remaining transparent connective tissue underneath the skin. The superficial musculature should now be exposed; displace this with a foam surgical spear.</li>
	<li>Use foam surgical spears (or a curette) to clear away the remaining, deeper musculature of the target vertebra (thoracic 12). To create a seat for the backplate, also clear away muscle from the posterior aspect of thoracic 11, and the anterior aspect of thoracic 13. Control any bleeding by applying gentle pressure with a surgical spear or use a minimal pulse with a cautery gun. Continue to remove the remaining muscle away from the tendons using forceps</li>
	<li>Once muscles are removed, carefully detach the tendons by cutting with forceps. There should be plenty of space for visualizing and manipulating the cord when this step is complete. Check that the dura matter of the inter-vertebral space, the semi-transparent laminar bone, the central superficial blood vessel underneath the bone, and anterior radiating artery are now clearly visible.</li>
	<li>Wet the region with warm aCSF. Use the microdrill to repetitively thin the laminar bone using straight strokes parallel to the long axis of the spinal cord (Figure 2, Figure 3). If desired, utilize a gliding stage to rotate the surgical platform for enhanced ergonomic comfort (for example, a right-handed operator may rotate the surgical platform counterclockwise for the drilling step). 
	Note: The gliding stage used is constructed of an upper aluminum plate that slides &#177;15 mm with respect to the fixed base plate.</li>
	<li>Gently grasp the superficial spinous process with forceps and lift the vertebra; the bone should lift away easily. If there is resistance, repeat bone thinning with the drill and if necessary use iris scissors, being careful to aim the scissor tips upward to avoid damaging the tissue. 
	Note: In order to maintain the dura intact, it is essential not to tug on the bone.</li>
	<li>Use #4 forceps to clear away any bone shards. Use a surgical spear to apply gentle steady pressure to control any bleeding. Rinse tissue with warm aCSF. Do not allow the tissue to dry out.</li>
</ol>
5. Cover-glass Implantation
<ol>
	<li>Gently apply a 3 mm borosilicate cover-glass to the exposed cord.</li>
	<li>Ensure agarose is cooled to 39 &#176;C. Using a small spatula, apply warm 2% agarose/aCSF to the edge of the cover-glass and allow capillary action to draw it under the surface. 
	Note: At temperatures below 39 &#176;C, the agarose may start to gel. If this occurs, rewarm using a water bath or microwave. Some operators prefer to first apply one drop of agarose and lay the cover-glass on top.</li>
	<li>Apply tissue adhesive to the exposed bony articular processes of the intact adjacent vertebra at thoracic level 11 vertebral spine and thoracic level 13 vertebral spine. Apply additional tissue adhesive in a ring around the laminectomy site, over the adjacent tendon and transverse process. 
	Note: Tissue adhesive is required for proper adherence of the dental cement in the subsequent steps. The articular processes form a natural seat on which the backplate can rest stably (Figure 3). Adhesion to the articular processes will form the strongest points of attachment.</li>
	<li>Mix dental cement with accelerant in a porcelain mixing tray. Utilize a small spatula to transfer dental cement onto the tissue adhesive layer. Use dental cement to adhere the backplate to the surgical field, centered over the window. Allow 10 minutes for the dental cement to cure. 
	Note: The firm adhesion of the backplate to the anterior and posterior articular processes provides the fundamental structural stability of the implant.</li>
	<li>Use additional dental cement to fill in the interior base of the backplate, and the underside of the backplate : tissue interface. 
	Note: The extra application of dental cement improves adherence and reduces risk of leakage of microscope objective immersion fluid (saline) out the bottom of the backplate.</li>
	<li>Advance the forked backplate holder to the appropriate position over the window. Secure the backplate into the backplate holder with screws. 
	Note: This protocol utilized a custom-machined backplate holder (Figure 2G&#8210;H).</li>
	<li>Apply saline to the backplate to test for leakage. If any fluid leaks, dry the area and apply more dental cement.</li>
</ol>
6. Imaging Preparation
<ol>
	<li>Transfer the animal on the surgical platform to the optical table. 
	Note: Our surgical platform, backplate holder, and isoflurane nose-cone holder can be transported between surgical and two-photon imaging stations as one unit, while applying continuous isoflurane anesthesia (Figure 2D,H). Similar units can be assembled from holding forks, beams, and supportive pillar posts obtainable from commercial sources (e.g. ThorLabs). For intravital microscopy, there should be at least 11 inches of clearance between the microscope objective and the optical table to accommodate the height of the surgical platform.</li>
	<li>Affix the surgical platform to the optical table using a stainless-steel mounting post and counterbored clamping fork.</li>
	<li>Apply fresh saline into the well of the backplate. Lower a water-immersion lens into the well.</li>
	<li>Use transmitted or epifluorescence light to identify the area of interest and focus. Switch to laser scanning mode and perform in vivo imaging according to the appropriate two-photon laser excitation wavelength, dichroics and bandpass filters for the fluorophores present in the tissue6.</li>
</ol>

<ol>
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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+procedure+for+implanting+a+spinal+chamber+for+longitudinal+in+vivo+imaging+of+the+mouse+spinal+cord.">A procedure for implanting a spinal chamber for longitudinal in vivo imaging of the mouse spinal cord.</a> Journal of Visualized Experiments. (94), (2014).</li><li>Fenrich, K. K., Weber, P., Rougon, G., Debarbieux, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Implanting+glass+spinal+cord+windows+in+adult+mice+with+experimental+autoimmune+encephalomyelitis.">Implanting glass spinal cord windows in adult mice with experimental autoimmune encephalomyelitis.</a> Journal of Visualized Experiments. (82), e50826 (2013).</li><li>Figley, S. 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=A+spinal+cord+window+chamber+model+for+in+vivo+longitudinal+multimodal+optical+and+acoustic+imaging+in+a+murine+model.">A spinal cord window chamber model for in vivo longitudinal multimodal optical and acoustic imaging in a murine model.</a> PLOS ONE. 8 (3), e58081 (2013).</li><li>Helmchen, F., Denk, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deep+tissue+two-photon+microscopy.">Deep tissue two-photon microscopy.</a> Nature Methods. 2 (12), 932-940 (2005).</li><li>Liebner, 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=Functional+morphology+of+the+blood-brain+barrier+in+health+and+disease.">Functional morphology of the blood-brain barrier in health and disease.</a> Acta Neuropathologica. 135 (3), 311-336 (2018).</li><li>Shen, L., Weber, C. R., Turner, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+tight+junction+protein+complex+undergoes+rapid+and+continuous+molecular+remodeling+at+steady+state.">The tight junction protein complex undergoes rapid and continuous molecular remodeling at steady state.</a> Journal of Cell Biology. 181 (4), 683-695 (2008).</li><li>Knowland, D., 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=Stepwise+recruitment+of+transcellular+and+paracellular+pathways+underlies+blood-brain+barrier+breakdown+in+stroke.">Stepwise recruitment of transcellular and paracellular pathways underlies blood-brain barrier breakdown in stroke.</a> Neuron. 82, 1-15 (2014).</li><li>Lutz, S. 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=Caveolin1+Is+Required+for+Th1+Cell+Infiltration,+but+Not+Tight+Junction+Remodeling,+at+the+Blood-Brain+Barrier+in+Autoimmune+Neuroinflammation.">Caveolin1 Is Required for Th1 Cell Infiltration, but Not Tight Junction Remodeling, at the Blood-Brain Barrier in Autoimmune Neuroinflammation.</a> Cell Reports. 21 (8), 2104-2117 (2017).</li><li>Harrison, 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=Vertebral+landmarks+for+the+identification+of+spinal+cord+segments+in+the+mouse.">Vertebral landmarks for the identification of spinal cord segments in the mouse.</a> Neuroimage. 68, 22-29 (2013).</li><li>Cupido, A., Catalin, B., Steffens, H., Kirchhoff, F., Bakota, L., Brandt, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Laser Scanning Microscopy and Quantitative Image Analysis of Neuronal Tissue. , 37-50 (2014).</li><li>Sekiguchi, K. 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The authors have nothing to disclose.

The method described here allows for stable imaging of the spinal cord in mice through a glass window. This method has been applied to assess BBB remodeling in transgenic eGFP:Claudin5+/- mice that express a fluorescent BBB tight junction protein, but it could be applied equally well for studies of any fluorescent proteins or cells in the spinal cord.
Multiple methods for laminectomy and spinal cord stabilization have been developed. All protocols address stabilizing the spinal cord during ima...

Transgenic animal models expressing fluorescent proteins, when combined with intravital microscopy, provide a powerful platform for addressing biology and pathophysiology. To apply these techniques to the spinal cord, specialized protocols are required to prepare the spinal cord for imaging. One such strategy is to conduct a laminectomy and spinal cord window implantation. The key features of an ideal laminectomy protocol for microscopy include preservation of native tissue structure and function, stability of the imaging field, quick processing time, and reproducibility of results. A particular challenge is to stabilize the imaging field against the motion induced by respiration and heartbeat. Multiple ex vivo and in vivo strategies have been reported to achieve these goals1,2,3,4,5. Most in vivo methods involve clamping the sides of the spinal column2,4 and is often followed by implanting a rigid metal apparatus3,4 for stability during surgery and downstream imaging applications. Clamping the spinal column can potentially compromise blood flow and induce blood-brain barrier (BBB) protein remodeling.
The purpose of this method is to make the intact spinal cord available for optical imaging in the living mouse while minimizing the invasiveness of the protocol and improving outcomes. We describe a single laminectomy and cover-glass implantation procedure paired with a minimally invasive oval plastic 3D-printed backplate that still achieves robust mechanical stability. The backplate is directly adhered to the anterior and posterior vertebral spines with dental cement. The backplate is equipped with lateral extension arms with screw holes that rigidly attach to the microscope stage via a metal arm. This effectively anchors the intact anterior and posterior vertebra to the microscope stage, providing mechanical resistance to the motion artifact that would otherwise be introduced by respiration and heartbeat. The method has been optimized for laminectomy of a single vertebra at thoracic level 12, omitting the clamps utilized in alternative strategies for stability during in vivo imaging. The procedure is rapid, taking approximately 30 min per mouse.
This protocol can be used to study disease mechanisms of the BBB. The BBB is a dynamic microvascular system comprised of endothelial cells, vascular smooth muscle, pericytes, and astrocyte foot processes that provide a highly selective environment for the central nervous system (CNS). Representative data depict the application of this protocol in transgenic mice engineered to express enhanced green fluorescent protein (eGFP):Claudin-5, a BBB tight junction protein. The provided backplate printing files can also be customized for alternative applications.

<table>
	<tbody>
		<tr>
			<td>3D printer</td>
			<td>Raise3D</td>
			<td>Pro2</td>
			<td>For printing backplates</td>
		</tr>
		<tr>
			<td>PLA 3D printing filament</td>
			<td>Inland</td>
			<td>PLA+-175-B</td>
			<td>Black plastic 3D printing material</td>
		</tr>
		<tr>
			<td>3D CAD software</td>
			<td>Dassault Systemes</td>
			<td>Solidworks software</td>
			<td>used to design 3D shapes</td>
		</tr>
		<tr>
			<td>3D printer software</td>
			<td>Raise3D</td>
			<td>Ideamaker software</td>
			<td>software used to interface with the 3D printer</td>
		</tr>
		<tr>
			<td>3D printed oval backplate</td>
			<td>custom</td>
			<td></td>
			<td>Stabilizing imaging field</td>
		</tr>
		<tr>
			<td>Surgical dissecting microscope</td>
			<td>Leica</td>
			<td>M205 C</td>
			<td>Equipped with Leica FusionOptics, Planapo 0.63x M-series objective, and gliding stage</td>
		</tr>
		<tr>
			<td>Microscope camera</td>
			<td>Leica</td>
			<td>MC170</td>
			<td>HD color camera for visualizing surgical field</td>
		</tr>
		<tr>
			<td>Gliding stage</td>
			<td>Leica</td>
			<td>10446301</td>
			<td>The gliding stage is constructed of two metal plates. The base plate is fixed. The upper plate slides on greased interface to allow rotational and linear movement.</td>
		</tr>
		<tr>
			<td>Surgical station and stabilization fork</td>
			<td>Whale Manufactoring</td>
			<td>custom</td>
			<td>Laminectomy</td>
		</tr>
		<tr>
			<td>SomnoSuite low-flow isoflurane delivery unit</td>
			<td>Kent Scientific</td>
			<td>SS-01</td>
			<td>Surgical anesthesia administration with integrated digitial vaporizer</td>
		</tr>
		<tr>
			<td>Stainless steel 1.5 inch mounting post</td>
			<td>ThorLabs</td>
			<td>P50/M</td>
			<td>For mounting surgical station onto optical table for two-photon imaging</td>
		</tr>
		<tr>
			<td>Counterbored Clamping Fork for 1.5&#34; mounting Post</td>
			<td>ThorLabs</td>
			<td>PF175</td>
			<td>For stabilizing surgical station mount onto optical table for two-photon imaging</td>
		</tr>
		<tr>
			<td>Ideal bone microdrill</td>
			<td>Harvard apparatus</td>
			<td>72-6065</td>
			<td>Thinning bone for laminectomy</td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>Fisher Scientific</td>
			<td>15-462-10</td>
			<td>Warming saline</td>
		</tr>
		<tr>
			<td>Cautery gun</td>
			<td>FST</td>
			<td>18010-00</td>
			<td>Cauterizing minor bleeds</td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>Benchmark</td>
			<td>BF11222</td>
			<td>1.9&#8221; x 4.5&#8221; silicone heater with 20&#8221; Teflon leads, 10W, 5V</td>
		</tr>
		<tr>
			<td>K type thermocoupled rectal probe</td>
			<td>Physitemp</td>
			<td>RET3</td>
			<td>Measuring mouse body temperature</td>
		</tr>
		<tr>
			<td>petroleum jelly</td>
			<td>Sigma</td>
			<td>8009-03-8</td>
			<td>Lubricating rectal probe</td>
		</tr>
		<tr>
			<td>Feedback-regulated thermal controller</td>
			<td>custom</td>
			<td>NA</td>
			<td>Commercially available alternatives include the Physitemp TCAT series</td>
		</tr>
		<tr>
			<td>PVA Surgical eye spears</td>
			<td>Beaver-visitec international</td>
			<td>40400-8</td>
			<td>Absorbing blood</td>
		</tr>
		<tr>
			<td>Electric trimmer</td>
			<td>Wahl</td>
			<td>41590-0438</td>
			<td>Trimming mouse fur</td>
		</tr>
		<tr>
			<td>Blade, #11</td>
			<td>FST</td>
			<td>14002-14</td>
			<td>Surgical tool</td>
		</tr>
		<tr>
			<td>Forceps, #5</td>
			<td>FST</td>
			<td>11254-20</td>
			<td>Surgical tool</td>
		</tr>
		<tr>
			<td>Forceps, #4</td>
			<td>FST</td>
			<td>14002-14</td>
			<td>Surgical tool</td>
		</tr>
		<tr>
			<td>Titatnium toothed forceps</td>
			<td>WPI</td>
			<td>555047FT</td>
			<td>Surgical tool</td>
		</tr>
		<tr>
			<td>Titanium Iris scissors</td>
			<td>WPI</td>
			<td>555562S</td>
			<td>Surgical tool</td>
		</tr>
		<tr>
			<td>Vetbond tissue adhesive</td>
			<td>3M</td>
			<td>084-1469SB</td>
			<td>Preparing tissue surface for dental acrylic</td>
		</tr>
		<tr>
			<td>Ceramic mixing tray</td>
			<td>Jack Richeson</td>
			<td>420716</td>
			<td>Mixing dental acrylic agent with accelerant</td>
		</tr>
		<tr>
			<td>Orthojet dental acrylic</td>
			<td>Lang Dental</td>
			<td>1520BLK, 1503BLK</td>
			<td>Permanently bonding backplate to tissue</td>
		</tr>
		<tr>
			<td>Small round cover glass, #1 thickness, 3 mm</td>
			<td>Harvard apparatus</td>
			<td>64-0720</td>
			<td>optical window</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Fisher Scientific</td>
			<td>7647-14-5</td>
			<td>For aCSF</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fisher Scientific</td>
			<td>7447-40-7</td>
			<td>For aCSF</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Fisher Scientific</td>
			<td>50-99-7</td>
			<td>For aCSF</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>7365-45-9</td>
			<td>For aCSF</td>
		</tr>
		<tr>
			<td>MgCl2&#183;6H2O</td>
			<td>Fisher Scientific</td>
			<td>7791-18-6</td>
			<td>For aCSF</td>
		</tr>
		<tr>
			<td>CaCl2&#183;2H2O</td>
			<td>Fisher Scientific</td>
			<td>10035-04-8</td>
			<td>For aCSF</td>
		</tr>
		<tr>
			<td>Carprofen</td>
			<td>Rimadyl</td>
			<td>QM01AE91</td>
			<td>Analgesia</td>
		</tr>
		<tr>
			<td>Bacteriostatic water</td>
			<td>Henry Schein</td>
			<td>2587428</td>
			<td>Diluent for carprofen</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Henry Schein</td>
			<td>11695-6776-2</td>
			<td>Anesthesia</td>
		</tr>
		<tr>
			<td>Lactated ringer solution</td>
			<td>Baxter</td>
			<td>0338-0117-04</td>
			<td>Hydration for mouse</td>
		</tr>
		<tr>
			<td>Agarose High EEO</td>
			<td>Sigma</td>
			<td>A9793</td>
			<td>gel point 34-37 degrees C</td>
		</tr>
		<tr>
			<td>Opthalmic lubricating ointment</td>
			<td>Akwa Tears</td>
			<td>68788-0697</td>
			<td>Prevent corneal drying</td>
		</tr>
		<tr>
			<td>MOM Two-Photon Microscope</td>
			<td>Sutter</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Implanted glass windows and intravital two-photon microscopy provides a useful tool for assessing dynamic changes in CNS proteins. The functional integrity of the BBB is influenced by the expression, subcellular localization, and turnover rates of tight junction proteins7. Previous studies have demonstrated that tight junction proteins undergo rapid and dynamic remodeling at steady state8. The currently described laminectomy and glass window...

Watch this Scientific Journal Video about Laminectomy and Spinal Cord Window Implantation in the Mouse at JoVE.com

Laminectomy and Spinal Cord Window Implantation in the Mouse

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 ...

laminectomy-and-spinal-cord-window-implantation-in-the-mouse

Research

JoVE Journal

Neuroscience

11.9K Views.  University of Illinois at Chicago College of Medicine. This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

Video: Laminectomy and Spinal Cord Window Implantation in the Mouse

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Diffusion Imaging in the Rat Cervical Spinal Cord

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a diagnostic and prognostic tool in cases of disease or injury. Diffusion weighted imaging (DWI), is sensitive to the thermal motion of water molecules and allows for inferences of tissue microstructure. This report describes a protocol to acquire and analyze DWI of the rat cervical spinal cord on a small-bore animal system. It demonstrates an imaging setup for the live anesthetized animal and recommends a DWI acquisition protocol for high-quality imaging, which includes stabilization of the cord and control of respiratory motion. Measurements with diffusion weighting along different directions and magnitudes (b-values) are used. Finally, several mathematical models of the resulting signal are used to derive maps of the diffusion processes within the spinal cord tissue that provide insight into the normal cord and can be used to monitor injury or disease processes noninvasively.

The goal of this protocol is to obtain high-quality diffusion weighted magnetic resonance imaging (DWI) of the rat spinal cord for noninvasive characterization of tissue microstructure. This protocol describes optimizations of the MRI sequence, radiofrequency coil, and analysis methods to enable DWI images free from artifacts.

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

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

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

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

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

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

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...