Name: in vivo imaging of cerebrospinal fluid transport through the intact mouse skull using fluorescence macroscopy
Uploaded: 2019-07-29T13:00:00.000+00:00
Duration: 6 min 22 s
Description: Watch this Scientific Journal Video about In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy at JoVE.com

This work was funded by the National Institute of Neurological Disorders and Stroke and the National Institute on Aging (US National Institutes of Health; R01NS100366 and RF1AG057575 to MN), the Fondation Leducq Transatlantic Networks of Excellence Program, and the EU Horizon 2020 research and innovation programme (grant no. 666881; SVDs@target). We would also like to thank Dan Xue for expert assistance with graphic illustrations.

All experiments were approved by the University Committee on Animal Resources (UCAR, Protocol No. 2011-023) at the University of Rochester and performed according to the NIH Guide for the Care and Use of Laboratory Animals.
1. Preparing the cisterna magna cannula, head plate, and head holder
<ol>
	<li>Sterilize all surgical instruments and head plates before surgery. 
	NOTE: Fluorescent tracers are delivered directly into CSF via a cisterna magna cannulation. For detailed instructions on this procedure, please refer to Xavier et al9.</li>
	<li>Briefly, using a needle driver, break the tip of a 30G x 12.7 mm (&#189; inch) needle, &#190; of the way down, and place the blunt end of the needle into one end of polyethylene 10 (PE10) tubing (about 45 cm long). Make sure that only the bevel is protruding out of the border of the PE10 tubing.</li>
	<li>Break off &#188; of the beveled end of another 30G needle and place the blunt end of the remaining needle into the other end of the PE10 tubing, with the plastic Luer-lock still attached.</li>
	<li>Fill a 100 &#181;L glass syringe with sterile, artificial CSF (aCSF: 126 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 2 mM MgSO4, 2 mM CaCl2, 10 mM glucose, and 26 mM NaHCO3).</li>
	<li>Attach the syringe to the end of the line and fill it with aCSF until it reaches the tip of the beveled needle. It is important that the syringe is backfilled with aCSF and not air, as an air column is more prone to variable infusion volumes.</li>
	<li>For chronic experiments, leave the line filled with aCSF and skip Step 1.7.</li>
	<li>For acute experiments, place the syringe on an infusion pump and withdraw approximately 5 mm of air, which will prevent mixing. Afterwards, withdraw the total volume of tracer(s) that is desired for the experiment (20% extra is recommended due to dead space losses). 
	NOTE: The PE10 tubing should be long enough so that the air bubble does not enter the plastic cuff of the glass syringe when loading the tracer. For a typical experiment, the protocol uses 10 &#181;L of bovine serum albumin conjugated to Alexa Fluor 647 (BSA-647) diluted in aCSF at 0.5%.</li>
	<li>Confirm that the head plate fits into the head holder and that it is the right size for the mouse being used. Anatomical markers to ensure this are: the top border of the window aligns with the interocular line and the posterior border falls rostral to the occipital crest. 
	NOTE: Stainless-steel head plates can be sterilized and reused. Most cyanoacrylate mixtures can be removed with an acetone solution.</li>
</ol>
2. Surgical procedure
<ol>
	<li>Weigh and anesthetize the mouse (e.g. ketamine/xylazine; 100 mg/kg ketamine, 10 mg/kg xylazine; i.p.).</li>
	<li>Once the mouse no longer responds to a toe pinch, moisten the neck and head with sterile water and shave using clippers. Once the area is shaved, wipe the area with an alcohol swab again to remove any residual hair. 
	NOTE: Moistening the fur before clipping dramatically reduces the amount of hair in the imaging window after the incision.</li>
	<li>Place the mouse in a stereotactic frame on top of a temperature-controlled pad and apply petroleum ophthalmic ointment to the mouse&#8217;s eyes to ensure they do not dry out.</li>
	<li>Clean the exposed skin with a chlorhexidine swab. After 2 min, remove the chlorhexidine with an alcohol wipe. Finally, apply an iodine solution which can be left to dry.</li>
	<li>Inject subcutaneous analgesia (0.25% Bupivacaine HCl) to the top of the skull and the neck.</li>
	<li>Starting on the part of the neck that covers the occipital crest, make a midline cut in the overlying skin and continue rostrally towards the interorbital line. Incise laterally to the border where the temporal muscle inserts into the skull. Remove all of the skin of the fusiform incision to expose both the frontal and parietal bones. 
	CAUTION: The retroorbital sinus lies caudal to the eyes and large branches of the posterior facial vein lie rostral to the pinna. Be careful when making the incision to spare these structures. If this occurs, stop bleeding by maintaining hemostasis with a sterile cotton swab for several minutes and continue.</li>
	<li>Irrigate the skull with sterile saline and clean the surface using cotton swabs so that it is free of debris and hair since these will interfere with the image quality. Skull transparency is best preserved by leaving the periosteum and overlying fascia intact. 
	NOTE: If these structures are accidentally removed, the skull can become dry and opaque over time. Re-moisten with aCSF or use a mixture of paraffin oil and glycerol to reduce skull reflectance in acute experiments21.</li>
	<li>Proceed to inserting the cisterna magna cannula - for surgical details of this procedure, refer to Xavier et al9.</li>
	<li>After inserting the CM cannula, apply a mixture of dental cement with cyanoacrylate glue to the ventral side of the head plate around the border and place it on the skull so that the anterior border of the head plate aligns with the posterior tip of the nasal bone and the posterior border aligns with the anterior aspect of the interparietal bone, making sure that the sagittal suture (midline) is centered and straight relative to the window (Figure 1B).</li>
	<li>Fix the position of the head plate using a couple drops of glue accelerator. Fill any remaining gaps with the cement mixture and cure it with accelerator. 
	CAUTION: Make sure the cyanoacrylate does not come in contact with the mouse&#8217;s eyes or block the imaging window.</li>
	<li>Glue the CM cannula to the head plate to ensure that these do not become detached during transport or when the mouse wakes up.</li>
	<li>For acute experiments, skip to Step 2.16.</li>
	<li>For chronic experiments, apply a thin layer of clear-drying cyanoacrylate glue to the exposed skull, being careful not to create bubbles as these will interfere with the imaging. The glue provides protection for the skull and will not interfere with the imaging. Make sure that the glue covers the exposed skull all the way to the skin at the incision border.</li>
	<li>Use a hemostat clamp to hold the aCSF-filled PE10 tubing 2-3 cm from the CM and cut the line with a high-temperature cautery tip. Once it is separated, flatten the melted PE10 tubing to seal the cannula. 
	CAUTION: Make sure not to release the clamp until after the cannula has been sealed and confirm there are no leaks in the tubing to prevent a CSF fistula.</li>
	<li>Administer carprofen (5 mg/kg every 24 h for 3 days; i.p. or s.c.) and return the mouse to a temperature-controlled, single-housed cage and allow it to recover for at least 24 h prior to imaging. Do not leave the mouse unattended until it regains sufficient consciousness to maintain sternal recumbency. Intact cranial windows remain stable for several weeks. 
	NOTE: The chronic experiment can be paused here. 
	CAUTION: Some postoperative complications are: cephalohematoma/subgaleal hematomas, CSF fistula, and infection.</li>
	<li>For acute experiments, place the mouse into the head holder using the head plate so the skull is in a fixed position. Make sure to check anesthesia level and place a heating pad under the mouse. The mouse is now ready to be taken to the macroscope. 
	CAUTION: Carefully transport the mouse together with the infusion pump on a cart. If the CM cannulation becomes dislodged, it will cause a drop in intracranial pressure and any CSF leaks will alter the experimental results.</li>
</ol>
3. Preparing the mouse for imaging
NOTE: The protocol varies depending on whether the imaging experiment will be performed on an anesthetized (start at Step 3.1) or awake (start at Step 3.2) mouse.
<ol>
	<li>Anesthetized Mice
	<ol>
		<li>Place the head holder on the stage of the macroscope, making sure there are no kinks in the line from the syringe pump to the CM cannula and that it is not taut since this could affect the tracer infusion.</li>
		<li>Observe respiratory rate and pink coloration of the mucous membranes, indicating good oxygenation. Inject saline subcutaneously to secure hydration level if necessary. Check to make sure the animal is sufficiently anesthetized and re-dose if needed.</li>
		<li>Turn on the macroscope camera and LED and start LIVE mode.</li>
		<li>Confirm the magnification of the imaging field so that the nasofrontal suture at the top of the field and the lambdoid suture at the bottom can clearly be visualized, with the sagittal suture parallel and centered to the middle of the image (Figure 1C). Once in place, secure the head holder to the macroscope stage with tape.</li>
		<li>Focus the macroscope on the exposed skull. Despite macroscopes having a relatively large depth of field, the curvature of the mouse&#8217;s skull only allows for a specific area to be in focus. Best results are obtained when the focal plane is located on the lateral sides of the parietal bones posterior to the coronal sutures. 
		NOTE: This is the location of the middle cerebral arteries (MCA), where most CSF inflow occurs (Figure 1D,E)10.</li>
	</ol>
	</li>
	<li>Awake Mice
	<ol>
		<li>Prior to the imaging session, let the animals recover for at least 24 h from the head plate surgery. Longer recovery periods (5-7 days) are also recommended. During this time, train the mouse to be head fixed on the stage in the restraint tube for 0.5-1 h per day for the duration of the recovery period. 
		NOTE: Habituation allows the mouse to be attached to the head holder without anesthesia and reduces stress and anxiety during the actual experiment. If this is not feasible and anesthesia does not interfere with the experiment, an induction dose of an inhaled anesthetic (e.g., isoflurane 2% at 1-2 L/min O2 flow rate) can be used to quickly attach the mouse to the head stage.</li>
		<li>Once the mouse is fixed to the head holder and in the restraint tube, follow Steps 3.1.1-3.1.5.</li>
	</ol>
	</li>
</ol>
4. Infusion of fluorescent CSF tracers
<ol>
	<li>Acute CM Cannulation
	<ol>
		<li>Since the tracer was already loaded into the cannula before being placed in the cisterna magna in Step 1.6, set the infusion pump to the desired rate and volume. Infusion paradigms routinely used are 5-10 &#181;L at 1-2 &#181;L/min, but these parameters can be adjusted depending on the particular experiment, or the size and age of the animal.</li>
	</ol>
	</li>
	<li>Chronic CM Cannulation
	<ol>
		<li>Prior to beginning the imaging session, follow Steps 1.2-1.7 for acute experiments to prepare the infusion line to deliver the tracers.</li>
		<li>Once the line is prepared, using a hemostat clamp cut the sealed end of the chronic CM cannula. Take the needle from the line prepared in Step 4.2.1 and gently insert it into the cannula. Release the clamp and using the syringe pump, advance the CSF tracer at the desired infusion rate (e.g., 2 &#181;L/min) for the experiment until the tracer reaches the implanted needle in the CM. 
		CAUTION: If the needle pierces through the tubing when connecting the line, remove the needle, cut the pierced segment and repeat this step. Any tear in the PE10 tubing will cause a leak and affect the results of the experiment.</li>
	</ol>
	</li>
</ol>
5. Setting up the Imaging Session
<ol>
	<li>Based on the fluorescent tracer being infused, determine the excitation wavelength and the exposure time for each channel. Choose the shortest exposure time necessary to visualize the tracer in order to maximize the temporal resolution of the time-lapse imaging. This exposure time will be used for all subsequent experiments aiming to compare CSF transport between different animals. 
	NOTE: One tip is to use the tracer in the CM line to adjust the exposure time. Although this is a useful first approach, this should be optimized over time.</li>
	<li>Choose the duration of the experiment and the intervals at which the images will be acquired. Experiments normally last between 30-60 min depending on what phase of glymphatic transport is of interest. Frame rates of 1 frame/min and faster are sufficient for most experiments.</li>
	<li>Set the file name and the saving directory.</li>
	<li>If the imaging will be collected in addition to simultaneous acquisition of other variables (e.g., electrocardiogram, arterial blood pressure, electrophysiology), the macroscope and pump can be programmed to be triggered with the data acquisition software. Check that the triggering function on the macroscope is correct before moving to the next step.</li>
</ol>
6. Transcranial optical imaging experiment
<ol>
	<li>Start the tracer infusion and the imaging at the same time.</li>
	<li>Routinely check the CM cannula and the PE10 tubing for any signs of a leak. If there is any tracer leaking at the CM, the results from this experiment must be excluded. 
	NOTE: If tracer starts to accumulate in the cerebellum and does not travel the glymphatic pathway of the MCA, it is likely that the CM cannula was injected into the cerebellum. This data should not be included.</li>
	<li>For acute experiments, after completion of the experiment, remove the mouse from the microscope and check that it is still adequately anesthetized. Quickly decapitate the mouse and harvest the brain tissue. Immersion-fix the brain in 4% paraformaldehyde (PFA) overnight at 4 &#176;C.
	<ol>
		<li>For chronic experiments, once the imaging is completed, remove the CM line from the cannula, flush with sterile aCSF and reseal the CM cannula with a high-temperature cautery tip. Remove the mouse from the head stand and return it to its cage. Repeat this process until further imaging is not required and then follow Step 6.3.</li>
	</ol>
	</li>
</ol>
7. Data analysis
NOTE: Matlab-based analyses, such as CSF front-tracking can extract large amounts of quantitative data from the tracer fronts in these imaging datasets10,22. However, these file types can also be easily imported and analyzed in open-source image analysis software like Fiji23.
<ol>
	<li>Import the image stacks into Fiji using the Bio-Formats import tool. This function will preserve the file metadata which includes the time and pixel resolution. Save the image stack as a .tiff file.</li>
	<li>Manually draw a region of interest (ROI) around the skull or the area of interest using the polygon selection tool. For example, in Figure 1G an ROI was drawn separately for the ipsilateral and for the contralateral hemisphere after a traumatic brain injury. Make sure to add the ROI into the ROI Manager (Analyze&#62;Tools&#62;ROI Manager) and save it. 
	NOTE: CSF transport can be quantified in two main ways: 1) mean fluorescence intensity over time or 2) influx area expressed as a percentage of the ROI or total area (mm2) after having applied a threshold. The latter method (Figure 1H) will be described in the next steps.</li>
	<li>Set the threshold on the frame with maximal fluorescence (Select Image&#62;Adjust&#62;Threshold&#8230;). Automated threshold methods such as Otsu are normally good at detecting CSF tracer. Apply the threshold.</li>
	<li>Before going forward, make sure that in Analyze&#62;Set Measurements, the options Area and Area Fraction are selected. Then in the ROI Manager, select More&#62;Multi Measure.</li>
	<li>Convert the %Area value into mm2 using the Area value of the ROI from the output. The %Area values reflect the percent of the dorsal cortical surface with CSF tracer.</li>
	<li>Plot CSF tracer influx in mm2 as a function of time.</li>
</ol>

<ol>
	<li>Tumani, H., Huss, A., Bachhuber, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+cerebrospinal+fluid+and+barriers+-+anatomic+and+physiologic+considerations.">The cerebrospinal fluid and barriers - anatomic and physiologic considerations.</a> Handbook of Clinical Neurology. , 21-32 (2017).</li><li>Jessen, N. A., Munk, A. S., Lundgaard, I., Nedergaard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Glymphatic+System:+A+Beginner's+Guide.">The Glymphatic System: A Beginner's Guide.</a> Neurochemical Research. 40 (12), 2583-2599 (2015).</li><li>Iliff, J. J., 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=Impairment+of+glymphatic+pathway+function+promotes+tau+pathology+after+traumatic+brain+injury.">Impairment of glymphatic pathway function promotes tau pathology after traumatic brain injury.</a> The Journal of Neuroscience. 34 (49), 16180-16193 (2014).</li><li>Iliff, J. J., 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+paravascular+pathway+facilitates+CSF+flow+through+the+brain+parenchyma+and+the+clearance+of+interstitial+solutes,+including+amyloid+beta.">A paravascular pathway facilitates CSF flow through the brain parenchyma and the clearance of interstitial solutes, including amyloid beta.</a> Science Translational Medicine. 4 (147), (2012).</li><li>Louveau, 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=Structural+and+functional+features+of+central+nervous+system+lymphatic+vessels.">Structural and functional features of central nervous system lymphatic vessels.</a> Nature. 523 (7560), 337-341 (2015).</li><li>Peng, W., 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=Suppression+of+glymphatic+fluid+transport+in+a+mouse+model+of+Alzheimer's+disease.">Suppression of glymphatic fluid transport in a mouse model of Alzheimer's disease.</a> Neurobiology of Disease. 93, 215-225 (2016).</li><li>Da Mesquita, ., 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+aspects+of+meningeal+lymphatics+in+ageing+and+Alzheimer's+disease.">Functional aspects of meningeal lymphatics in ageing and Alzheimer's disease.</a> Nature. 560 (7717), 185-191 (2018).</li><li>Gaberel, T., 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=Impaired+glymphatic+perfusion+after+strokes+revealed+by+contrast-enhanced+MRI:+a+new+target+for+fibrinolysis.">Impaired glymphatic perfusion after strokes revealed by contrast-enhanced MRI: a new target for fibrinolysis.</a> Stroke. 45 (10), 3092-3096 (2014).</li><li>Xavier, A. L. R., 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=Cannula+Implantation+into+the+Cisterna+Magna+of+Rodents.">Cannula Implantation into the Cisterna Magna of Rodents.</a> Journal of Visualized Experiments. 10 (135), (2018).</li><li>Plog, B. 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=Transcranial+optical+imaging+reveals+a+pathway+for+optimizing+the+delivery+of+immunotherapeutics+to+the+brain.">Transcranial optical imaging reveals a pathway for optimizing the delivery of immunotherapeutics to the brain.</a> JCI Insight. 3 (23), (2018).</li><li>Kress, B. T., 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=Impairment+of+paravascular+clearance+pathways+in+the+aging+brain.">Impairment of paravascular clearance pathways in the aging brain.</a> Annals of Neurology. 76 (6), 845-861 (2014).</li><li>Mestre, H., 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=Flow+of+cerebrospinal+fluid+is+driven+by+arterial+pulsations+and+is+reduced+in+hypertension.">Flow of cerebrospinal fluid is driven by arterial pulsations and is reduced in hypertension.</a> Nature Communications. 9 (1), 4878 (2018).</li><li>Xie, 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=Sleep+drives+metabolite+clearance+from+the+adult+brain.">Sleep drives metabolite clearance from the adult brain.</a> Science. 342 (6156), 373-377 (2013).</li><li>Plog, B. A., Nedergaard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Glymphatic+System.">The Glymphatic System.</a> in Central Nervous System Health and Disease: Past, Present, and Future. Annual Review of Pathology. 13, 379-394 (2018).</li><li>Iliff, J. J., 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=Brain-wide+pathway+for+waste+clearance+captured+by+contrast-enhanced+MRI.">Brain-wide pathway for waste clearance captured by contrast-enhanced MRI.</a> Journal of Clinical Investigation. 123 (3), 1299-1309 (2013).</li><li>Davoodi-Bojd, 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=Modeling+glymphatic+system+of+the+brain+using+MRI.">Modeling glymphatic system of the brain using MRI.</a> Neuroimage. 188, 616-627 (2019).</li><li>Lee, H., 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=The+Effect+of+Body+Posture+on+Brain+Glymphatic+Transport.">The Effect of Body Posture on Brain Glymphatic Transport.</a> The Journal of Neuroscience. 35 (31), 11034-11044 (2015).</li><li>Hablitz, L. 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=Increased+glymphatic+influx+is+correlated+with+high+EEG+delta+power+and+low+heart+rate+in+mice+under+anesthesia.">Increased glymphatic influx is correlated with high EEG delta power and low heart rate in mice under anesthesia.</a> Science Advances. 5 (2), (2019).</li><li>Rasmussen, M. K., Mestre, H., Nedergaard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+glymphatic+pathway+in+neurological+disorders.">The glymphatic pathway in neurological disorders.</a> The Lancet Neurology. 17 (11), 1016-1024 (2018).</li><li>Mestre, H., 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=Aquaporin-4-dependent+glymphatic+solute+transport+in+the+rodent+brain.">Aquaporin-4-dependent glymphatic solute transport in the rodent brain.</a> Elife. 7, (2018).</li><li>Monai, H., 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=Calcium+imaging+reveals+glial+involvement+in+transcranial+direct+current+stimulation-induced+plasticity+in+mouse+brain.">Calcium imaging reveals glial involvement in transcranial direct current stimulation-induced plasticity in mouse brain.</a> Nature Communications. 7, 11100 (2016).</li><li>Munk, 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=PDGF-B+Is+Required+for+Development+of+the+Glymphatic+System.">PDGF-B Is Required for Development of the Glymphatic System.</a> Cell Reports. 26 (11), 2955-2969 (2019).</li><li>Schindelin, J., 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=Fiji:+an+open-source+platform+for+biological-image+analysis.">Fiji: an open-source platform for biological-image analysis.</a> Nature Methods. 9 (7), 676-682 (2012).</li><li>Ren, Z., 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=Hit+&amp;+Run'+model+of+closed-skull+traumatic+brain+injury+(TBI)+reveals+complex+patterns+of+post-traumatic+AQP4+dysregulation.">Hit &amp; Run' model of closed-skull traumatic brain injury (TBI) reveals complex patterns of post-traumatic AQP4 dysregulation.</a> Journal of Cerebral Blood Flow &amp; Metabolism. 33 (6), 834-845 (2013).</li><li>Plog, B. 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=Biomarkers+of+traumatic+injury+are+transported+from+brain+to+blood+via+the+glymphatic+system.">Biomarkers of traumatic injury are transported from brain to blood via the glymphatic system.</a> The Journal of Neuroscience. 35 (2), 518-526 (2015).</li><li>Ma, Q., Ineichen, B. V., Detmar, M., Proulx, S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Outflow+of+cerebrospinal+fluid+is+predominantly+through+lymphatic+vessels+and+is+reduced+in+aged+mice.">Outflow of cerebrospinal fluid is predominantly through lymphatic vessels and is reduced in aged mice.</a> Nature Communications. 8 (1), 1434 (2017).</li><li>Roth, T. 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=Transcranial+amelioration+of+inflammation+and+cell+death+after+brain+injury.">Transcranial amelioration of inflammation and cell death after brain injury.</a> Nature. 505 (7482), 223-228 (2014).</li><li>Xu, H. T., Pan, F., Yang, G., Gan, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Choice+of+cranial+window+type+for+in+vivo+imaging+affects+dendritic+spine+turnover+in+the+cortex.">Choice of cranial window type for in vivo imaging affects dendritic spine turnover in the cortex.</a> Nature Neuroscience. 10 (5), 549-551 (2007).</li><li>Ma, Q., 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=Rapid+lymphatic+efflux+limits+cerebrospinal+fluid+flow+to+the+brain.">Rapid lymphatic efflux limits cerebrospinal fluid flow to the brain.</a> Acta Neuropathologica. 137 (1), 151-165 (2019).</li><li>Silasi, G., Xiao, D., Vanni, M. P., Chen, A. C., Murphy, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intact+skull+chronic+windows+for+mesoscopic+wide-field+imaging+in+awake+mice.">Intact skull chronic windows for mesoscopic wide-field imaging in awake mice.</a> Journal of Neuroscience Methods. 267, 141-149 (2016).</li></ol>

The authors have nothing to disclose.

We have described a detailed protocol for performing transcranial CSF imaging in live mice using commercially-available fluorescent macroscopes and tracers. This technique is simple and minimally-invasive, yet quantitative. In vivo imaging correlates well with sensitive methods such as liquid scintillation counting of radio-labeled tracers including 3H-dextran and 14C-inulin after CM delivery, and with ex vivo coronal section quantification10,18</...

Cerebrospinal fluid (CSF) bathes the brain and spinal cord and is involved in maintaining homeostasis, supplying nutrients, and regulating intracranial pressure1. CSF in the subarachnoid space enters the brain through a network of perivascular spaces (PVS) surrounding cortical pial arteries and then flows down along penetrating arterioles2. Once in the parenchyma, CSF exchanges with interstitial fluid (ISF), carrying harmful metabolites such as amyloid-&#946; (A&#946;) and tau protein aggregates out of the brain through low resistance white matter tracts and perivenous spaces2,3. This pathway is dependent on astroglial aquaporin-4 (AQP4) channels and has therefore been termed the glial-lymphatic (glymphatic) system4. Waste products of the neuropil are ultimately cleared from the CSF-ISF through lymphatic vessels near cranial nerves and in the meninges out towards the cervical lymph nodes5. The failure of this system has been implicated in several neurologic diseases such as Alzheimer&#8217;s disease6,7, traumatic brain injury3, and ischemic and hemorrhagic stroke8.&#160;
CSF transport can be visualized by infusing tracers into the cisterna magna (CM)9,10 and glymphatic studies in the past have mainly utilized two-photon microscopy4,11,12,13, magnetic resonance imaging (MRI)14,15,16,17, and ex vivo imaging3,6,11,18 to evaluate tracer kinetics. Two-photon microscopy is a suitable method for detailed imaging of CSF tracers in PVSs and the parenchyma due to its high spatial resolution, however, it has a narrow field of view and requires an invasive cranial window or thinning of the skull. Ex vivo imaging, in combination with immunohistochemistry, enables multilevel analyses ranging from single cells up to the whole brain19. However, the process of perfusion-fixation that is required to observe the post-mortem tissue produces profound changes in CSF flow direction and collapses the PVS, significantly altering the distribution and the location of the tracers12. Finally, while MRI can track CSF flow throughout the entire murine and human brain, it lacks spatial and temporal resolution of perivascular flow.
A new technique, transcranial macroscopic imaging, solves some of these limitations by enabling wide-field imaging of perivascular CSF transport in the entire dorsal cortex of living mice. This type of imaging is done with an epifluorescent macroscope using a multiband filter cube, tunable LED light source, and high-efficiency CMOS camera10. These set-ups are able to resolve PVSs up to 1-2 mm below the skull surface and can detect fluorophores up to 5-6 mm below the cortical surface while leaving the skull entirely intact10. Multiband filters and LEDs that can quickly tune the excitation wavelength enable the use of multiple fluorophores allowing CSF to be labeled with tracers of different molecular weights and chemical properties in the same experiment.
This procedure requires a simple, minimally invasive surgery to expose the skull and place a light-weight head plate to stabilize the head during the imaging session. Tracers can be delivered into the CM without drilling into the skull or penetrating the cortical tissue with pipettes or cannulas9,20. Both CM cannulas and head plates remain stable for several days to weeks and facilitate more complex experimental designs compared to the classical end-point visualization. This protocol describes how transcranial macroscopic imaging is used to study glymphatic system function following acute or chronic injection of fluorescent CSF tracer into the CM of anesthetized/sleeping or awake mice.

<table><tbody><tr><td>0.25% Bupivacaine HCl</td><td></td><td></td><td>University of Rochester Vivarium</td></tr><tr><td>100 &amp;micro;L Gastight Syringe Model 1710 TLL, PTFE Luer Lock</td><td>Hamilton Company</td><td>81020</td><td></td></tr><tr><td>A-M Systems Dental Cement Powder</td><td>Fisher Scientific</td><td>NC9991371</td><td></td></tr><tr><td>Carprofen</td><td></td><td></td><td>University of Rochester Vivarium</td></tr><tr><td>Chlorhexidine</td><td>Prevantics</td><td>B10800</td><td></td></tr><tr><td>CMOS Camera</td><td>Hammamatsu</td><td>ORCA Flash 4.0</td><td></td></tr><tr><td>Head Plate</td><td>University of Rochester</td><td>No catalog #</td><td>Custom made at the machine shop at the University of Rochester</td></tr><tr><td>High-Temperature Cautery</td><td>Bovie Medical Corporation</td><td>AA01</td><td></td></tr><tr><td>Insta-set Accelerator</td><td>Bob Smith Industries</td><td>BSI-151</td><td></td></tr><tr><td>Isoflurane - Fluriso</td><td>Vet One</td><td>502017</td><td>University of Rochester Vivarium</td></tr><tr><td>Ketamine</td><td></td><td></td><td>Strong Memorial Hospital Pharmacy</td></tr><tr><td>Krazy Glue</td><td>Elmer&#039;s Products, Inc</td><td>No catalog #, see link in comments</td><td>https://www.amazon.com/Krazy-Glue-KG48348MR-Advance-Multicolor/dp/B000BKO6DG</td></tr><tr><td>Micropore Surgical tape</td><td>Fisher Scientific</td><td>19-027-761</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-aldrich</td><td>P6148</td><td></td></tr><tr><td>PE10 - Polyethylene .011&quot; x .024&quot; per ft., 100 ft. continuous</td><td>Braintree Scientific</td><td>PE10 100 FT</td><td></td></tr><tr><td>Pump 11 Elite Infusion Only Dual Syringe</td><td>Harvard Apparatus</td><td>70-4501</td><td></td></tr><tr><td>PURALUBE VET OINTMENT</td><td>Dechra</td><td></td><td></td></tr><tr><td>Puritan PurSwab Cotton Tipped Cleaning Sticks</td><td>Fisher Scientific</td><td>22-029-553</td><td></td></tr><tr><td>Research Macro Zoom Microscope</td><td>Olympus</td><td>MVX10</td><td></td></tr><tr><td>Simple Head Holder Plate (for mice)</td><td>Narishige International USA Inc</td><td>MAG-1</td><td></td></tr><tr><td>Single-use Needles, BD Medical</td><td>VWR</td><td>BD305106</td><td></td></tr><tr><td>Sterile Alcohol Prep Pads</td><td>Fisher Scientific</td><td>22-363-750</td><td></td></tr><tr><td>Tunable LED</td><td>PRIOR Lumen 1600-LED</td><td></td><td></td></tr><tr><td>Xylazine</td><td></td><td></td><td>University of Rochester Vivarium</td></tr></tbody></table>

in vivo imaging of cerebrospinal fluid transport through the intact mouse skull using fluorescence macroscopy

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and magnetic resonance imaging (MRI) have enabled in vivo quantification of CSF flow but they are limited by reduced imaging volumes and low spatial resolution, respectively. Recent work has found that CSF enters the brain parenchyma through a network of perivascular spaces surrounding the pial and penetrating arteries of the rodent cortex. This perivascular entry of CSF is a primary driver of the glymphatic system, a pathway implicated in the clearance of toxic metabolic solutes (e.g., amyloid-&#946;). Here, we illustrate a new macroscopic imaging technique that allows real-time, mesoscopic imaging of fluorescent CSF tracers through the intact skull of live mice. This minimally-invasive method facilitates a multitude of experimental designs and enables single or repeated testing of CSF dynamics. Macroscopes have high spatial and temporal resolution and their large gantry and working distance allow for imaging while performing tasks on behavioral devices. This imaging approach has been validated using two-photon imaging and fluorescence measurements obtained from this technique strongly correlate with ex vivo fluorescence and quantification of radio-labeled tracers. In this protocol, we describe how transcranial macroscopic imaging can be used to evaluate glymphatic transport in live mice, offering an accessible alternative to more costly imaging modalities.

CSF influx is imaged on an epifluorescent macroscope (Figure 1A), which allows for mesoscopic imaging of CSF tracer transport in the murine cortex. The whole-skull head plate permits the visualization of the rostral nasal bones, both frontal and parietal bones in the center, and the rostral portion of the interparietal bone caudally (Figure 1B). During imaging, the nasofrontal, sagittal, coronal, and lambdoid sutures can be readily identified (<strong class="xfi...

Watch this Scientific Journal Video about In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy at JoVE.com

In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

Transcranial optical imaging allows wide-field imaging of cerebrospinal fluid transport in the cortex of live mice through an intact skull.

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and ...

in-vivo-imaging-cerebrospinal-fluid-transport-through-intact-mouse

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

13.6K Views.  University of Rochester Medical Center. Transcranial optical imaging allows wide-field imaging of cerebrospinal fluid transport in the cortex of live mice through an intact skull.

Video: In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

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 ...

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, inflammatory and metabolic conditions, using in vivo fluorescence microscopy. A closed cranial window is placed over the left parieto-occipital cortex of the mice. Local microcirculation is recorded in real time through the window using epi and fluorescence illumination, and measurements of vessels diameters and red blood cell (RBC) velocities are performed. RBC velocity is measured using real-time cross-correlation and/or fluorescent-labeled erythrocytes. Leukocyte and platelet adherence to pial vessels and assessment of perfusion and vascular leakage are made with the help of fluorescence-labeled markers such as Albumin-FITC and anti-CD45-TxR antibodies. Microcirculation can be repeatedly video-recorded over several days. We used for the first time the close window brain intravital microscopy to study the pial microcirculation to follow dynamic changes during the course of Plasmodium berghei ANKA infection in mice and show that expression of CM is associated with microcirculatory dysfunctions characterized by vasoconstriction, profound decrease in blood flow and eventually vascular collapse.

Intravital microscopy to follow temporal and spatial hemodynamic and inflammatory events in the pial microcirculation.

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, ...

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 ...

Combining Intravital Fluorescent Microscopy (IVFM) with Genetic Models to Study Engraftment Dynamics of Hematopoietic Cells to Bone Marrow Niches

Increasing evidence indicates that normal hematopoiesis is regulated by distinct microenvironmental cues in the BM, which include specialized cellular niches modulating critical hematopoietic stem cell (HSC) functions1,2. Indeed, a more detailed picture of the hematopoietic microenvironment is now emerging, in which the endosteal and the endothelial niches form functional units for the regulation of normal HSC and their progeny3,4,5. New studies have revealed the importance of perivascular cells, adipocytes and neuronal cells in maintaining and regulating HSC function6,7,8. Furthermore, there is evidence that cells from different lineages, i.e. myeloid and lymphoid cells, home and reside in specific niches within the BM microenvironment. However, a complete mapping of the BM microenvironment and its occupants is still in progress.
Transgenic mouse strains expressing lineage specific fluorescent markers or mice genetically engineered to lack selected molecules in specific cells of the BM niche are now available. Knock-out and lineage tracking models, in combination with transplantation approaches, provide the opportunity to refine the knowledge on the role of specific &#34;niche&#34; cells for defined hematopoietic populations, such as HSC, B-cells, T-cells, myeloid cells and erythroid cells. This strategy can be further potentiated by merging the use of two-photon microscopy of the calvarium. By providing in vivo high resolution imaging and 3-D rendering of the BM calvarium, we can now determine precisely the location where specific hematopoietic subsets home in the BM and evaluate the kinetics of their expansion over time. Here, Lys-GFP transgenic mice (marking myeloid cells)9 and RBPJ knock-out mice (lacking canonical Notch signaling)10 are used in combination with IVFM to determine the engraftment of myeloid cells to a Notch defective BM microenvironment.

Combining Intravital Fluorescent Microscopy IVFM with Genetic Models to Study Engraftment Dynamics of Hematopoietic Cells to Bone Marrow Niches

Intravital fluorescence microscopy (IVFM) of the calvarium is applied in combination with genetic animal models to study the homing and engraftment of hematopoietic cells into bone marrow (BM) niches.

Increasing evidence indicates that normal hematopoiesis is regulated by distinct microenvironmental cues in the BM, which include specialized cellular ...

Developmental Biology

In Vivo Optical Imaging of Brain Tumors and Arthritis Using Fluorescent SapC-DOPS Nanovesicles

We describe a multi-angle rotational optical imaging (MAROI) system for in vivo monitoring of physiopathological processes labeled with a fluorescent marker. Mouse models (brain tumor and arthritis) were used to evaluate the usefulness of this method. Saposin C (SapC)-dioleoylphosphatidylserine (DOPS) nanovesicles tagged with CellVue Maroon (CVM) fluorophore were administered intravenously. Animals were then placed in the rotational holder (MARS) of the in vivo imaging system. Images were acquired in 10&#176; steps over 380&#176;. A rectangular&#160;region of interest (ROI) was placed across the full image width at the model disease site. Within the ROI, and for every image, mean fluorescence intensity was computed after background subtraction. In the mouse models studied, the labeled nanovesicles were taken up in both the orthotopic and transgenic brain tumors, and in the arthritic sites (toes and ankles). Curve analysis of the multi angle image ROIs determined the angle with the highest signal. Thus, the optimal angle for imaging each disease site was characterized. The MAROI method applied to imaging of fluorescent compounds is a noninvasive, economical, and precise tool for in vivo quantitative analysis of the disease states in the described mouse models.

In Vivo Optical Imaging of Brain Tumors and Arthritis Using Fluorescent SapC-DOPS Nanovesicles

We describe a multi-angle rotational optical imaging (MAROI) system for in vivo quantitation of a fluorescent marker delivered by saposin C (SapC)-dioleoylphosphatidylserine (DOPS) nanovesicles. Employing mouse models of cancer and arthritis, we demonstrate how the MAROI signal curve analysis can be used for the precise mapping and biological characterization of disease processes.

We describe a multi-angle rotational optical imaging (MAROI) system for in vivo monitoring of physiopathological processes labeled with a fluorescent ...

Medicine

Quantification of Cerebral Vascular Architecture using Two-photon Microscopy in a Mouse Model of HIV-induced Neuroinflammation

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic neuroinflammatory state within the central nervous system (CNS), thought to be driven principally by virally-mediated activation of microglia and brain resident macrophages. HIV-1 infection is also accompanied by changes in cerebrovascular blood flow (CBF), raising the possibility that HIV-associated chronic neuroinflammation may lead to changes in CBF and/or in cerebral vascular architecture. To address this question, we have used a mouse model for HIV-induced neuroinflammation, and we have tested whether long-term exposure to this inflammatory environment may damage brain vasculature and result in rarefaction of capillary networks. In this paper we describe a method to quantify changes in cortical capillary density in a mouse model of neuroinflammatory disease (HIV-1 Tat transgenic mice). This generalizable approach employs in vivo two-photon imaging of cortical capillaries through a thin-skull cortical window, as well as ex vivo two-photon imaging of cortical capillaries in mouse brain sections. These procedures produce images and z-stack files of capillary networks, respectively, which can be then subjected to quantitative analysis in order to assess changes in cerebral vascular architecture.

This paper describes a method by which the vascular architecture in the brain can be quantified using in vivo and ex vivo two-photon microscopy.

Human Immunodeficiency Virus 1 (HIV-1) infection frequently results in HIV-1 Associated Neurocognitive Disorders (HAND), and is characterized by a chronic ...

Precise Brain Mapping to Perform Repetitive In Vivo Imaging of Neuro-Immune Dynamics in Mice

The central nervous system (CNS) is regulated by a complex interplay of neuronal, glial, stromal, and vascular cells that facilitate its proper function. Although studying these cells in isolation in vitro or together ex vivo provides useful physiological information; salient features of neural cell physiology will be missed in such contexts. Therefore, there is a need for studying neural cells in their native in vivo environment. The protocol detailed here describes repetitive in vivo two-photon imaging of neural cells in the rodent cortex as a tool to visualize and study specific cells over extended periods of time from hours to months. We describe in detail the use of the grossly stable brain vasculature as a coarse map or fluorescently labeled dendrites as a fine map of select brain regions of interest. Using these maps as a visual key, we show how neural cells can be precisely relocated for subsequent repetitive in vivo imaging. Using examples of in vivo imaging of fluorescently-labeled microglia, neurons, and NG2+ cells, this protocol demonstrates the ability of this technique to allow repetitive visualization of cellular dynamics in the same brain location over extended time periods, that can further aid in understanding the structural and functional responses of these cells in normal physiology or following pathological insults. Where necessary, this approach can be coupled to functional imaging of neural cells, e.g., with calcium imaging. This approach is especially a powerful technique to visualize the physical interaction between different cell types of the CNS in vivo when genetic mouse models or specific dyes with distinct fluorescent tags to label the cells of interest are available.

This protocol describes a chronic cranial window implantation technique that can be used for longitudinal imaging of neuro-glio-vascular structures, interactions, and function in both healthy and diseased conditions. It serves as a complementary alternative to the transcranial imaging approach that, while often preferred, possesses some critical limitations.

The central nervous system (CNS) is regulated by a complex interplay of neuronal, glial, stromal, and vascular cells that facilitate its proper function. ...

Immunology and Infection

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.

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.

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

Intravital Imaging of Fluorescent Protein Expression in Mice with a Closed-Skull Traumatic Brain Injury and Cranial Window Using a Two-Photon Microscope

The goal of this protocol is to demonstrate how to longitudinally visualize the expression and localization of a protein of interest within specific cell types of an animal's brain, upon exposure to exogenous stimuli. Here, the administration of a closed-skull traumatic brain injury (TBI) and simultaneous implantation of a cranial window for subsequent longitudinal intravital imaging in mice is shown. Mice are intracranially injected with an adeno-associated virus (AAV) expressing enhanced green fluorescent protein (EGFP) under a neuronal specific promoter. After 2 to 4 weeks, the mice are subjected to a repetitive TBI using a weight drop device over the AAV injection location. Within the same surgical session, the mice are implanted with a metal headpost and then a glass cranial window over the TBI impacting site. The expression and cellular localization of EGFP is examined using a two-photon microscope in the same brain region exposed to trauma over the course of months.

This study demonstrates delivery of a repetitive traumatic brain injury to mice and simultaneous implantation of a cranial window for subsequent intravital imaging of a neuron-expressed EGFP using two-photon microscopy.

The goal of this protocol is to demonstrate how to longitudinally visualize the expression and localization of a protein of interest within specific cell ...

Fluorescence Imaging of Neuro-Immune Dynamics Using a Skull Implant

<a href='https://appjove.unicartagenaproxy.elogim.com/v/61454/precise-brain-mapping-to-perform-repetitive-vivo-imaging-neuro-immune'>Source:&#160; Bisht, K., et al. Precise brain mapping to perform repetitive in vivo imaging of neuro-immune dynamics in mice. J. Vis. Exp. (2020)</a>This video demonstrates fluorescence imaging of neuro-immune dynamics in a transgenic mouse with a skull implant. Vascular and neuronal stability is imaged alongside dynamic microglial changes, reflecting immune responses to neuronal activity.

Source:&#160; Bisht, K., et al. Precise brain mapping to perform repetitive in vivo imaging of neuro-immune dynamics in mice. J. Vis. Exp. (2020)This ...

In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

Summary

Explore More Videos

Chapters in this video

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

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

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

Combining Intravital Fluorescent Microscopy (IVFM) with Genetic Models to Study Engraftment Dynamics of Hematopoietic Cells to Bone Marrow Niches

In Vivo Optical Imaging of Brain Tumors and Arthritis Using Fluorescent SapC-DOPS Nanovesicles

Quantification of Cerebral Vascular Architecture using Two-photon Microscopy in a Mouse Model of HIV-induced Neuroinflammation

Precise Brain Mapping to Perform Repetitive In Vivo Imaging of Neuro-Immune Dynamics in Mice

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

Intravital Imaging of Fluorescent Protein Expression in Mice with a Closed-Skull Traumatic Brain Injury and Cranial Window Using a Two-Photon Microscope

Fluorescence Imaging of Neuro-Immune Dynamics Using a Skull Implant