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>

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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.

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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

Research

JoVE Journal

Neuroscience

13.5K 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

Chronic Imaging of Mouse Visual Cortex Using a Thinned-skull Preparation

In vivo imaging using two-photon laser scanning microscopy (2PLSM) allows the study of living cells and neuronal processes in the intact brain. The technique presented here allows the imaging of the same area of the brain at several time points (chronic imaging) with microscopic resolution allowing the tracking of dendritic spines which are the small structures that represent the majority of postsynaptic excitatory sites in the CNS. The ability to clearly resolve fine cortical structures over several time points has many advantages, specifically in the study of brain plasticity in which morphological changes at synapses and circuit remodeling may help explain underlying mechanisms. In this video and supplementary material, we show a protocol for chronic in vivo imaging of the intact brain using a thinned-skull preparation. The thinned-skull preparation is a minimally invasive approach, which avoids potential damage to the dura and/or cortex, thus reducing the onset of an inflammatory response. When this protocol is performed correctly, it is possible to clearly monitor changes in dendritic spine characteristics in the intact brain over a prolonged period of time.

In this video and supplemental material, we show a protocol for chronic in vivo imaging of the intact brain using a thinned-skull preparation.

In vivo imaging using two-photon laser scanning microscopy (2PLSM) allows the study of living cells and neuronal processes in the intact brain. The ...

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 ...

Isolation of Cerebrospinal Fluid from Rodent Embryos for use with Dissected Cerebral Cortical Explants

The CSF is a complex fluid with a dynamically varying proteome throughout development and in adulthood. During embryonic development, the nascent CSF differentiates from the amniotic fluid upon closure of the anterior neural tube. CSF volume then increases over subsequent days as the neuroepithelial progenitor cells lining the ventricles and the choroid plexus generate CSF. The embryonic CSF contacts the apical, ventricular surface of the neural stem cells of the developing brain and spinal cord. CSF provides crucial fluid pressure for the expansion of the developing brain and distributes important growth promoting factors to neural progenitor cells in a temporally-specific manner. To investigate the function of the CSF, it is important to isolate pure samples of embryonic CSF without contamination from blood or the developing telencephalic tissue. Here, we describe a technique to isolate relatively pure samples of ventricular embryonic CSF that can be used for a wide range of experimental assays including mass spectrometry, protein electrophoresis, and cell and primary explant culture. We demonstrate how to dissect and culture cortical explants on porous polycarbonate membranes in order to grow developing cortical tissue with reduced volumes of media or CSF. With this method, experiments can be performed using CSF from varying ages or conditions to investigate the biological activity of the CSF proteome on target cells.

The ventricular cerebrospinal fluid (CSF) bathes the neuroepithelial and cerebral cortical progenitor cells during early brain development in the embryo. Here we describe the method developed to isolate ventricular CSF from rodent embryos of different ages in order to investigate its biological function. In addition, we demonstrate our cerebral cortical explant dissection and culture technique that allows for explant growth with minimal volumes of culture medium or CSF.

The CSF is a complex fluid with a dynamically varying proteome throughout development and in adulthood. During embryonic development, the nascent CSF ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then radial migration in the molecular layer and the Purkinje cell layer to reach the internal granular layer of the cerebellar cortex. Default in migratory processes induces either cell death or misplacement of the neurons, leading to deficits in diverse cerebellar functions. Centripetal granule cell migration involves several mechanisms, such as chemotaxis and extracellular matrix degradation, to guide the cells towards their final position, but the factors that regulate cell migration in each cortical layer are only partially known. In our method, acute cerebellar slices are prepared from P10 rats, granule cells are labeled with a fluorescent cytoplasmic marker and tissues are cultured on membrane inserts from 4 to 10 hr before starting real-time monitoring of cell migration by confocal macroscopy at 37 &#176;C in the presence of CO2. During their migration in the different cortical layers of the cerebellum, granule cells can be exposed to neuropeptide agonists or antagonists, protease inhibitors, blockers of intracellular effectors or even toxic substances such as alcohol or methylmercury to investigate their possible role in the regulation of neuronal migration.

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal cerebellum development, immature granule cells originating from the germinal zone exhibit distinct modalities of migration to reach their final destination and to establish neuronal networks. This protocol describes the preparation of cerebellar slices and the confocal macroscopic approach used to investigate the factors that regulate neuronal migration.

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then ...

Sample Preparation for Endopeptidomic Analysis in Human Cerebrospinal Fluid

This protocol describes a method developed to identify endogenous peptides in human cerebrospinal fluid (CSF). For this purpose, a previously developed method based on molecular weight cut-off (MWCO) filtration and mass spectrometric analysis was combined with an offline high-pH reverse phase HPLC pre-fractionation step.
Secretion into CSF is the main pathway for removal of molecules shed by cells of the central nervous system. Thus, many processes in the central nervous system are reflected in the CSF, rendering it a valuable diagnostic fluid. CSF has a complex composition, containing proteins that span a concentration range of 8 - 9 orders of magnitude. Besides proteins, previous studies have also demonstrated the presence of a large number of endogenous peptides. While less extensively studied than proteins, these may also hold potential interest as biomarkers.
Endogenous peptides were separated from the CSF protein content through MWCO filtration. By removing a majority of the protein content from the sample, it is possible to increase the sample volume studied and thereby also the total amount of the endogenous peptides. The complexity of the filtrated peptide mixture was addressed by including a reverse phase (RP) HPLC pre-fractionation step at alkaline pH prior to LC-MS analysis. The fractionation was combined with a simple concatenation scheme where 60 fractions were pooled into 12, analysis time consumption could thereby be reduced while still largely avoiding co-elution.
Automated peptide identification was performed by using three different peptide/protein identification software programs and subsequently combining the results. The different programs were complementary rather than comparable with less than 15% of the identifications overlapped between the three.

A method for mass spectrometric analysis of endogenous peptides in human cerebrospinal fluid (CSF) is presented. By employing molecular weight cut-off filtration, chromatographic pre-fractionation, mass spectrometric analysis and a subsequent combination of peptide identification strategies, it was possible to expand the known CSF peptidome nearly ten-fold compared to previous studies.

This protocol describes a method developed to identify endogenous peptides in human cerebrospinal fluid (CSF). For this purpose, a previously developed ...

In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters

This technique combines optical fiber mediated fluorescence recordings with the precise delivery of recombinant adeno-associated virus based gene reporters. This new and easy to use in vivo fluorescence monitoring system was developed to record the transcriptional rhythm of the clock gene, Cry1, in the suprachiasmatic nucleus (SCN) of freely moving mice. To do so, a Cry1 transcription fluorescence reporter was designed and packaged into Adeno-associated virus. Purified, concentrated virus was injected into the mouse SCN followed by the insertion of an optic fiber, which was then fixed onto the surface of the brain. The animals were returned to their home cages and allowed a 1-month post-operative recovery period to ensure sufficient reporter expression. Fluorescence was then recorded in freely moving mice via an in vivo monitoring system that was constructed at our institution. For the in vivo recording system, a 488 nm laser was coupled with a 1 &#215; 4 beam-splitter that divided the light into four laser excitation outputs of equal power. This setup enabled us to record from four animals simultaneously. Each of the emitted fluorescence signals was collected via a photomultiplier tube and a data acquisition card. In contrast to the previous bioluminescence in vivo circadian clock recording technique, this fluorescence in vivo recording system allowed the recording of circadian clock gene expression during the light cycle.

In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters

This newly developed fluorescence-based technology enables long-term monitoring of the transcription of circadian clock genes in the suprachiasmatic nucleus (SCN) of freely moving mice in real-time and at a high temporal resolution.

This technique combines optical fiber mediated fluorescence recordings with the precise delivery of recombinant adeno-associated virus based gene ...

An Improved Method for Collection of Cerebrospinal Fluid from Anesthetized Mice

The cerebrospinal fluid (CSF) is a valuable body fluid for analysis in neuroscience research. It is one of the fluids in closest contact with the central nervous system and thus, can be used to analyze the diseased state of the brain or spinal cord without directly accessing these tissues. However, in mice it is difficult to obtain from the cisterna magna due to its closeness to blood vessels, which often contaminate samples. The area for CSF collection in mice is also difficult to dissect to and often only small samples are obtained (maximum of 5-7 &#181;L or less). This protocol describes in detail a technique that improves on current methods of collection to minimize contamination from blood and allow for the abundant collection of CSF (on average 10-15 &#181;L can be collected). This technique can be used with other dissection methods for tissue collection from mice, as it does not impact any tissues during CSF extraction. Thus, the brain and spinal cord are not affected with this technique and remain intact. With greater CSF sample collection and purity, more analyses can be used with this fluid to further aid neuroscience research and better understand diseases affecting the brain and spinal cord.

This protocol describes an improved technique for the abundant collection of cerebrospinal fluid (CSF) with no contamination from blood. With greater sample collection and purity, more analyses can be performed using CSF to further our understanding of diseases that affect the brain and spinal cord.

The cerebrospinal fluid (CSF) is a valuable body fluid for analysis in neuroscience research. It is one of the fluids in closest contact with the central ...

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study of this movement in situ was only possible using radioisotopic pulse-labeling, which permits analysis of axonal transport in whole nerves with a temporal resolution of days and a spatial resolution of millimeters. To study neurofilament transport in situ with higher temporal and spatial resolution, we developed a hThy1-paGFP-NFM transgenic mouse that expresses neurofilament protein M tagged with photoactivatable GFP in neurons. Here we describe fluorescence photoactivation pulse-escape and pulse-spread methods to analyze neurofilament transport in single myelinated axons of tibial nerves from these mice ex vivo. Isolated nerve segments are maintained on the microscope stage by perfusion with oxygenated saline and imaged by spinning disk confocal fluorescence microscopy. Violet light is used to activate the fluorescence in a short axonal window. The fluorescence in the activated and flanking regions is analyzed over time, permitting the study of neurofilament transport with temporal and spatial resolution on the order of minutes&#160;and microns, respectively. Mathematical modeling can be used to extract kinetic parameters of neurofilament transport including the velocity, directional bias and pausing behavior from the resulting data. The pulse-escape and pulse-spread methods can also be adapted to visualize neurofilament transport in other nerves. With the development of additional transgenic mice, these methods could also be used to image and analyze the axonal transport of other cytoskeletal and cytosolic proteins in axons.

We describe fluorescence photoactivation methods to analyze the axonal transport of neurofilaments in single myelinated axons of peripheral nerves from transgenic mice that express a photoactivatable neurofilament protein.

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study ...