Authors would like to thank Charisse Montgomery for her editing service. A. Alomran&#8217;s work partially fulfills the requirements for a master&#8217;s degree in Pharmacology.This work is funded by The University of Toledo&#8217;s intramural startup fund for W.A.A and NIH grant (DK080640) for S.M.N.

The procedures for animal use were approved by the Institutional Animal Care and Use Committee (IACUC) of The University of Toledo in accordance with the guidelines of the Institutional Animal Care and Use Committee at the National Institutes of Health and the Guide for the Care and Use of Laboratory Animals.
1. Brain Extraction, Sectioning and Tissue Preparation
<ol>
	<li>Sacrifice wild-type mouse strain C57BL/6 by deeply euthanizing with CO2 asphyxiation for 5 min. Assure death by cervical dislocation.</li>
	<li>Clean the mouse head with 70% ethanol.</li>
	<li>Perform craniectomy using sterile scissors and forceps by first pulling the skin off, starting with the top of the head to expose the skull.</li>
	<li>Then, when the skull is exposed, remove the skull by peeling the bone piece-by-piece, starting from the posterior side and moving toward the anterior side. Be cautious not to destroy the brain ventricles.</li>
	<li>Collect the whole brain.</li>
	<li>Place the brain in a 100 mm Petri dish containing DMEM/High-Glucose supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin solution containing 10,000 units/ml of penicillin and 10,000 &#181;g/ml of streptomycin and pre-warmed to 37 &#176;C.</li>
	<li>Slice the brain on the median sagittal plane by hand with a sharp blade, and obtain the first 100-200 mm section from each half using a vibratome.</li>
	<li>Rinse the brain tissue with pre-warmed 37 &#176;C phosphate buffered saline (1x PBS) solution.</li>
	<li>Immediately place the brain section in DMEM/High-Glucose media pre-warmed to 37 &#176;C.</li>
</ol>
2. Live Imaging Configuration and Setup
<ol>
	<li>Place the brain tissue sections in 30 mm glass-bottom culture dishes containing 1 ml of DMEM/High-Glucose media. Adjust the microscope&#8217;s enclosed chamber environment to 37 &#176;C, 95%/5% O2/CO2 content (Figure 1).</li>
	<li>Using a 60X objective oil immersion lens, collect the ependymal cells/cilia images by first placing an oil drop on the 60X objective lens and focusing on the cells with regular DIC transmitted light.</li>
	<li>Then, follow the direction of the DMEM bubble movement as a guide to the location of the motile ependymal cilia as ciliary beatings create a kind of bubble movement in the area. Choose an area containing healthy cells with motile cilia in the brain&#8217;s lateral ventricle using the DIC filter. Once the ependymal cilia are found, adjust the light and focus to obtain a satisfactory image.</li>
	<li>Set the live imaging parameters according to a specific purpose using Metamorph imaging software. In the present demonstration, acquire twenty four-bit images with the camera binning set to 1 x 1 combined with 60X objective and 5-10 msec exposure time.</li>
	<li>Collect the DIC images by opening the microscope aperture to an optimal level in order to have a minimum exposure time. Observe live images stream to the camera to provide fast and immediate image acquisition without delay. Calculate the speed of cilia beating based on the requirement of the minimal exposure times to obtain sufficient image contrast.</li>
</ol>
3. Data Visualization and Analysis
<ol>
	<li>Calculate the number of cilia beatings by counting the number of beatings in 1 min. Do this by decreasing the speed of the video and counting the number of beats using a cell counter or similar tool.</li>
	<li>To calculate the frequency of beatings, multiply the exposure time at which the video is recorded by the number of the frames or time-lapse images acquired to get the number of sec. (Example: exposure time 5 msec 200 frames = 1,000 msec or 1 sec).</li>
	<li>Calculate the number of beatings in one second to obtain the frequency which is expressed as the number of beating per one sec. Do this by dividing the number of cilia beatings over a one-second time interval (Example: cilia beats 50 times in a 200 frames video recorded at exposure time of 5 msec i.e. 5 msec x 200 frames = 1 sec; now divide 50 beats by 1 sec = 50 Hz).</li>
	<li>Calculate the ciliary beating angle by evaluating the path taken by the ependymal cilia during both the power and recovery strokes. Perform this according to a previously described method, with minor modifications 11. The precise movement of individual cilia is observed during the complete beat cycle.</li>
	<li>On an acetate sheet placed over the monitor, draw a horizontal line along the ependymal edge and a vertical line through the midline position of the cilia at the start of the power stroke.</li>
	<li>Plot the precise position of the cilium frame by frame as it moves forward during the power stroke. In a similar manner, plot the movement of the cilia during the recovery stroke.</li>
	<li>Calculate the ciliary beating angle from the maximum deviation of the cilium from the midline during the power stroke as well as the recovery stroke.</li>
</ol>
4. Calcium Signal Recording
<ol>
	<li>After slicing the brain, briefly rinse the brain slice with 1x PBS or Dulbecco&#8217;s PBS (pH 7.0). Prepare fresh Fluo-2 to avoid fluorescence quenching and to obtain good signal-to-noise ratio.</li>
	<li>Prepare 1 mM stock solution of Fluo-2 solution in dimethylsulfoxide (DMSO), mix and vortex the solution for at least 5 min to ensure that Fluo-2 is homogenously dissolved in DMSO.</li>
	<li>Dilute the Fluo-2 stock solution in 500 ml of DMEM/High-Glucose supplemented with 2% B27 pre-warmed to 37 &#176;C to a final concentration of 20 mg/ml. 
	NOTE: B-27 is an optimized serum-free supplement containing vitamin A,&#160;antioxidant cocktail and&#160;insulin used to support short- or long-term viability of hippocampal and other CNS neurons.</li>
	<li>Immediately incubate the brain slice with 20 mg/ml Fluo-2 for 30 min at 37 &#176;C in a glass-bottom plate.</li>
	<li>To determine the optimal loading concentration of calcium fluorophore Fluo-2 and to avoid calcium dye cell toxicity, challenge cells with ATP, and check the cell viability by determining the time course and peak magnitude of calcium signals in response to ATP.</li>
	<li>Record the video for calcium oscillation at a capture rate of 5 msec for a minimum of 1 sec (200 frames per sec), with excitation and emission wavelengths of 488 nm and 515 nm, respectively (Movie 2).</li>
	<li>To distinguish the Fluo-2 calcium signal from autofluorescence or movement artifacts, ensure that the intensities emitted at 515 nm is monitored separately. 
	NOTE: Fluo-2 is not the calcium fluoremetric suitable to quantify intracellular calcium. However, it is an excellent dynamic calcium dye to detect fast calcium changes, such as calcium oscillations. For more accurate calcium quantification, Fura-2 is recommended. However, the dynamic changes of Fura-2 are limited by its ratiometric nature of the dye.</li>
	<li>Follow the formulas provided by the manufacturer to calculate the exact free intracellular calcium values. Example [Ca2+] = Kd &#215; (R&#8211;Rmin)/(Rmax &#8211; R). Where Kd is the dissociation constant of the dye from the released calcium, R is the measured fluorescence at 488, and Rmin and Rmax are the fluorescence ratios at minimum and maximum ion concentration 12.</li>
</ol>
5. Immunofluorescence Microscopy
<ol>
	<li>Fix the brain sections with phosphate buffered saline solution containing 4% paraformaldehyde (PFA) and 2% sucrose for 10 min. Alternatively, fix the whole brain with 4% PFA and then section into 50 mm sections using a cryostat.</li>
	<li>Wash the tissue three times with 1x PBS for 5 min each time.</li>
	<li>Incubate the brain slice with a solution of 0.1% Triton-X in 1x PBS for 5 min, then rinse three times with 1x PBS for 5 min each time.</li>
	<li>Incubate the brain slice with mouse primary antibody, antiacetylated a-tubulin, used at a dilution of 1:5,000 in 10% FBS in 1x PBS for one hour at room temperature (RT) or overnight at 4 &#176;C.</li>
	<li>Wash the tissue three times with 1x PBS for 5 min each time.</li>
	<li>Incubate the brain slice in secondary antibody, fluorescein anti-mouse IgG at a dilution of 1:500 in 10% FBS in 1x PBS solution for 1 hr at RT.</li>
	<li>Before observation under a fluorescent microscope, counterstain the section with 4&#39;,6-diamidino-2-phenylindole (DAPI) for 5 min to stain the nucleus (or DNA) 13. To minimize photobleaching, image the sections immediately with minimum exposure time possible.</li>
</ol>

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Ependymal cells along the lateral ventricle express functional p2x(7) receptors.</a> Purinergic signalling. 5 (3), 299-307 (2009).</li><li>Appelbe, O. K., 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=Disruption+of+the+mouse+jhy+gene+causes+abnormal+ciliary+microtubule+patterning+and+juvenile+hydrocephalus.">Disruption of the mouse jhy gene causes abnormal ciliary microtubule patterning and juvenile hydrocephalus.</a> Developmental biology. 382 (1), 172-185 (2013).</li><li>Sawamoto, K., 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=New+neurons+follow+the+flow+of+cerebrospinal+fluid+in+the+adult+brain+(New+York,+N.Y.).">New neurons follow the flow of cerebrospinal fluid in the adult brain (New York, N.Y.).</a> Science. 311 (5761), 629-632 (2006).</li><li>Banizs, B., 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=Dysfunctional+cilia+lead+to+altered+ependyma+and+choroid+plexus+function,+and+result+in+the+formation+of+hydrocephalus.">Dysfunctional cilia lead to altered ependyma and choroid plexus function, and result in the formation of hydrocephalus.</a> Development. 132 (23), 5329-5339 (2005).</li><li>Kawanabe, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cilostazol+prevents+endothelin-induced+smooth+muscle+constriction+and+proliferation.">Cilostazol prevents endothelin-induced smooth muscle constriction and proliferation.</a> PloS one. 7 (9), e44476 (2012).</li><li>Liu, T., Jin, X., Prasad, R. 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M., Jin, X., AbouAlaiwi, W. A., El-Jouni, W., Su, X., Zhou, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-motile+primary+cilia+as+fluid+shear+stress+mechanosensors.">Non-motile primary cilia as fluid shear stress mechanosensors.</a> Methods in enzymology. 525, 1-20 (2013).</li><li>AbouAlaiwi, W. 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=Survivin-induced+abnormal+ploidy+contributes+to+cystic+kidney+and+aneurysm+formation.">Survivin-induced abnormal ploidy contributes to cystic kidney and aneurysm formation.</a> Circulation. 129 (6), 660-672 (2014).</li><li>AbouAlaiwi, W. 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No conflicts of interest declared.

Described here is a protocol for the preparation of mouse brain tissue for both live-imaging and fluorescence microscopy that provides a rapid and sensitive close observation of the ependymal cilia within the brain ventricles. This technique is not restricted to the lateral ventricle only; it could be utilized to observe the cilia in other brain ventricles. This imaging technique provides a live stream that resembles the movement of the CSF by ciliary beating in an ex vivo setting. Moreover, it enables analysis ...

Cilia are sensory microtubule-based organelles that extend from the cell surface to the extracellular environment. Depending on their microtubule organization, cilia can be categorized into two types - &#8220;9+0&#8221; or &#8220;9+2&#8221;. Functionally, based on their motility, these can be classed as motile or non-motile cilia 1. Primary cilia is a term commonly used to denote &#8220;9+0&#8221; non-motile cilia. These have nine parallel doublet microtubules (denoted by &#8216;9&#8217;) and a central pair of microtubules is absent within the central sheath (denoted by &#8216;0&#8217;). However, some &#8220;9+0&#8221; cilia, such as nodal cilia, which regulate embryo laterality are motile 2. On the other hand, motile cilia are characterized, in addition to the nine parallel microtubule doublets, by an additional central pair of microtubule doublets and associated with dynein motor proteins to facilitate motility. In addition, some &#8220;9+2&#8221; cilia such as olfactory cilia are non-motile 3. Ependymal cells lining the brain ventricles and the central canal of the spinal cord are characterized by motile cilia that propel the cerebrospinal fluid (CSF) along the brain ventricles 4.
The overall goal of this method is to facilitate studying the motile cilia dynamics and structural abnormalities. The brain&#8217;s health and development heavily depend on efficient circulation of CSF within the brain ventricles. For instance, normal CSF flow and fluid balance require normal beating and functional ependymal cilia 5,6, which in turn play critical roles in regulating the directional movement of neuronal cells and stem cell migration 7. As such, abnormal ependymal cilia function or structure can lead to abnormal CSF flow, which is associated with hydrocephalus, a medical condition in which there is an abnormal accumulation of CSF in the ventricles of the brain. This may consequently cause increased intracranial pressure and progressive enlargement of the head, convulsion, tunnel vision, and mental disability 8.
The advantages of this technique over existing methods is that it allowed for the first time to report three distinct ependymal cell types: I, II, and III, based on their unique ciliary beating frequency and beating angle. These ependymal cells are localized within certain regions in the brain ventricles. Furthermore, the effects of age and pharmacological agents such as alcohol and cilostazol on altering the ependymal cell types or their localizations can be demonstrated, which was not possible before this classification of ependymal cells. Cilostazol is an inhibitor of phosphodiesterase-3, an enzyme that metabolizes cAMP to AMP and it also regulates intracellular calcium 9. Using high-speed fluorescence imaging analysis allows imaging and quantifying of the unique intracellular calcium oscillation properties of the ependymal cells. For instance, both alcohol and cilostazol significantly altered the ependymal ciliary beating frequency as well as the intracellular calcium oscillations properties, which in turn, could lead to a change in the cerebrospinal fluid volume replacement by ependymal cilia 10. In summary, this technique was key to provide the first evidence of three distinct types of ependymal cells with different calcium oscillation properties.
In the following section, a detailed step-by-step overview of the procedure is provided, paying close attention to tissue preparation and handling.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>DMEM/HIGH GLUCOSE</td>
			<td>Cellgro Mediatech Inc.</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Hyclone</td>
			<td>SH30088-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin</td>
			<td>Thermo Scientific</td>
			<td>SV30010</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline</td>
			<td>Thermo Scientific</td>
			<td>SH30256-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde&#160;</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710-SP</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich</td>
			<td>S-2395</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton-X</td>
			<td>Sigma-Aldrich</td>
			<td>T9284</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluo-2&#160;</td>
			<td>TEF Labs</td>
			<td>#0200</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Gibco</td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr>
			<td>VECTASHIELD HardSet Mounting Medium with DAPI</td>
			<td>Vector Labs</td>
			<td>H-1500</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-acetylated a-tubulin antibody</td>
			<td>Sigma-Aldrich</td>
			<td>T7451&#160;</td>
			<td>clone 6-11B1</td>
		</tr>
		<tr>
			<td>FITC Anti-mouse antibody</td>
			<td>Vector Labs</td>
			<td>FI-2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Culture plate</td>
			<td>VWR Vista Vision</td>
			<td>30-2041</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover Slip (18x18)</td>
			<td>VWR Vista Vision</td>
			<td>16004.326</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica Biosystems</td>
			<td>Leica VT1200S</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryostat</td>
			<td>Leica Biosystems</td>
			<td>Leica CM1860</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted Fluorescence Microscope</td>
			<td>Nikon&#160;</td>
			<td>Nikon TE2000</td>
			<td>60X oil&#160;</td>
		</tr>
		<tr>
			<td>Microscope cover glass 24x60mm</td>
			<td>VWR Vista Vision</td>
			<td>16004-312</td>
			<td></td>
		</tr>
		<tr>
			<td>Mounting Medium with DAPI</td>
			<td>Vector Laboratories</td>
			<td>H-1500</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI filter cube</td>
			<td>Chroma Technology</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

live imaging of the ependymal cilia in the lateral ventricles of the mouse brain

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various neurological deficits. The current ex vivo live imaging of motile ependymal cilia technique allows for a detailed study of ciliary dynamics following several steps. These steps include: mice euthanasia with carbon dioxide according to protocols of The University of Toledo&#8217;s Institutional Animal Care and Use Committee (IACUC); craniectomy followed by brain removal and sagittal brain dissection with a vibratome or sharp blade to obtain very thin sections through the brain lateral ventricles, where the ependymal cilia can be visualized. Incubation of the brain&#8217;s slices in a customized glass-bottom plate containing Dulbecco&#8217;s Modified Eagle&#8217;s Medium (DMEM)/High-Glucose at 37 &#176;C in the presence of 95%/5% O2/CO2 mixture is essential to keep the tissue alive during the experiment. A video of the cilia beating is then recorded using a high-resolution differential interference contrast microscope. The video is then analyzed frame by frame to calculate the ciliary beating frequency. This allows distinct classification of the ependymal cells into three categories or types based on their ciliary beating frequency and angle. Furthermore, this technique allows the use of high-speed fluorescence imaging analysis to characterize the unique intracellular calcium oscillation properties of ependymal cells as well as the effect of pharmacological agents on the calcium oscillations and the ciliary beating frequency. In addition, this technique is suitable for immunofluorescence imaging for ciliary structure and ciliary protein localization studies. This is particularly important in disease diagnosis and phenotype studies. The main limitation of the technique is attributed to the decrease in live motile cilia movement as the brain tissue starts to die.

Measuring ependymal cilia function in live mouse brain
The method described in this protocol is used to monitor ependymal cilia function and structure in the fresh tissue dissected from the mouse brain as well as to monitor and study cilia beating frequency. The steps followed to accomplish a complete experiment are depicted in a schematic flowchart (Figure 1). It is highly recommended that the experiment is conducted within a short time frame in order to keep the motile cilia...

Watch this Scientific Journal Video about Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain at JoVE.com

Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain

Using high-resolution differential interference contrast (DIC) microscopy, an ex vivo observation of the beating of motile ependymal cilia located within the mouse brain ventricles is demonstrated by live-imaging. The technique allows a recording of the unique ciliary beating frequency and beating angle as well as their intracellular calcium oscillation pacing properties.

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various ...

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13.6K Views.  University of Toledo, College of Pharmacy and Pharmaceutical Sciences. Using high-resolution differential interference contrast (DIC) microscopy, an ex vivo observation of the beating of motile ependymal cilia located within the mouse brain ventricles is demonstrated by live-imaging. The technique allows a recording of the unique ciliary beating frequency and beating angle as well as their intracellular calcium oscillation pacing properties.

Video: Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Manual Drainage of the Zebrafish Embryonic Brain Ventricles

Cerebrospinal fluid (CSF) is a protein rich fluid contained within the brain ventricles. It is present during early vertebrate embryonic development and persists throughout life. Adult CSF is thought to cushion the brain, remove waste, and carry secreted molecules1,2. In the adult and older embryo, the majority of CSF is made by the choroid plexus, a series of highly vascularized secretory regions located adjacent to the brain ventricles3-5. In zebrafish, the choroid plexus is fully formed at 144 hours post fertilization (hpf)6. Prior to this, in both zebrafish and other vertebrate embryos including mouse, a significant amount of embryonic CSF (eCSF) is present . These data and studies in chick suggest that the neuroepithelium is secretory early in development and may be the major source of eCSF prior to choroid plexus development7.
eCSF contains about three times more protein than adult CSF, suggesting that it may have an important role during development8,9. Studies in chick and mouse demonstrate that secreted factors in the eCSF, fluid pressure, or a combination of these, are important for neurogenesis, gene expression, cell proliferation, and cell survival in the neuroepithelium10-20. Proteomic analyses of human, rat, mouse, and chick eCSF have identified many proteins that may be necessary for CSF function. These include extracellular matrix components, apolipoproteins, osmotic pressure regulating proteins, and proteins involved in cell death and proliferation21-24. However, the complex functions of the eCSF are largely unknown.
We have developed a method for removing eCSF from zebrafish brain ventricles, thus allowing for identification of eCSF components and for analysis of the eCSF requirement during development. Although more eCSF can be collected from other vertebrate systems with larger embryos, eCSF can be collected from the earliest stages of zebrafish development, and under genetic or environmental conditions that lead to abnormal brain ventricle volume or morphology. Removal and collection of eCSF allows for mass spectrometric analysis, investigation of eCSF function, and reintroduction of select factors into the ventricles to assay their function. Thus the accessibility of the early zebrafish embryo allows for detailed analysis of eCSF function during development.

We present a method to collect cerebrospinal fluid (CSF) and to create a system which lacks CSF within the embryonic zebrafish brain ventricular system. This allows for further examination of CSF composition and its requirement during embryonic brain development.

Cerebrospinal fluid  (CSF) is a protein rich fluid contained within the brain ventricles. It is  present during early vertebrate embryonic development and ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Telomerase Activity in the Various Regions of Mouse Brain: Non-Radioactive Telomerase Repeat Amplification Protocol (TRAP) Assay

Telomerase, a ribonucleoprotein, is responsible for maintaining the telomere length and therefore promoting genomic integrity, proliferation, and lifespan. In addition, telomerase protects the mitochondria from oxidative stress and confers resistance to apoptosis, suggesting its possible importance for the surviving of non-mitotic, highly active cells such as neurons. We previously demonstrated the ability of novel telomerase activators to increase telomerase activity and expression in the various mouse brain regions and to protect motor neurons cells from oxidative stress. These results strengthen the notion that telomerase is involved in the protection of neurons from various lesions. To underline the role of telomerase in the brain, we here compare the activity of telomerase in male and female mouse brain and its dependence on age. TRAP assay is a standard method for detecting telomerase activity in various tissues or cell lines. Here we demonstrate the analysis of telomerase activity in three regions of the mouse brain by non-denaturing protein extraction using CHAPS lysis buffer followed by modification of the standard TRAP assay.
In this 2-step assay, endogenous telomerase elongates a specific telomerase substrate (TS primer) by adding TTAGGG 6 bp repeats (telomerase reaction). The telomerase reaction products are amplified by PCR reaction creating a DNA ladder of 6 bp increments. The analysis of the DNA ladder is made by 4.5% high resolution agarose gel electrophoresis followed by staining with highly sensitive nucleic acid stain.
Compared to the traditional TRAP assay that utilize 32P labeled radioactive dCTP&#39;s for DNA detection and polyacrylamide gel electrophoresis for resolving the DNA ladder, this protocol offers a non-toxic time saving TRAP assay for evaluating telomerase activity in the mouse brain, demonstrating the ability to detect differences in telomerase activity in the various female and male mouse brain region.

Telomerase Activity in the Various Regions of Mouse Brain: Non-Radioactive Telomerase Repeat Amplification Protocol TRAP Assay

Telomerase is expressed in the neonatal brain and also in distinct regions of the adult brain. We present a non-toxic time saving TRAP assay for the analysis of telomerase activity in various regions of the mouse brain and detection of differences in telomerase activity between male and female mouse brains.

Telomerase, a ribonucleoprotein, is responsible for maintaining the telomere length and therefore promoting genomic integrity, proliferation, and ...

3D Modeling of the Lateral Ventricles and Histological Characterization of Periventricular Tissue in Humans and Mouse

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional units of the ventricles are ciliated epithelial cells termed ependymal cells, which line the ventricles and through ciliary action are capable of generating laminar flow of CSF at the ventricle surface. This monolayer of ependymal cells also provides barrier and filtration functions that promote exchange between brain interstitial fluids (ISF) and circulating CSF. Biochemical changes in the brain are thereby reflected in the composition of the CSF and destruction of the ependyma can disrupt the delicate balance of CSF and ISF exchange. In humans there is a strong correlation between lateral ventricle expansion and aging. Age-associated ventriculomegaly can occur even in the absence of dementia or obstruction of CSF flow. The exact cause and progression of ventriculomegaly is often unknown; however, enlarged ventricles can show regional and, often, extensive loss of ependymal cell coverage with ventricle surface astrogliosis and associated periventricular edema replacing the functional ependymal cell monolayer. Using MRI scans together with postmortem human brain tissue, we describe how to prepare, image and compile 3D renderings of lateral ventricle volumes, calculate lateral ventricle volumes, and characterize periventricular tissue through immunohistochemical analysis of en face lateral ventricle wall tissue preparations. Corresponding analyses of mouse brain tissue are also presented supporting the use of mouse models as a means to evaluate changes to the lateral ventricles and periventricular tissue found in human aging and disease. Together, these protocols allow investigations into the cause and effect of ventriculomegaly and highlight techniques to study ventricular system health and its important barrier and filtration functions within the brain.

Using MRI scans (human), 3D imaging software, and immunohistological analysis, we document changes to the brain&#8217;s lateral ventricles. Longitudinal 3D mapping of lateral ventricle volume changes and characterization of periventricular cellular changes that occur in the human brain due to aging or disease are then modeled in mice.

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional ...

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

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

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

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

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or postmitotic neurons. Later, some progenitors switch to a gliogenic fate, adding to the astrocyte and oligodendrocyte populations. Using time-lapse video-microscopy of primary cerebral cortex cell cultures, it is possible to study the cellular and molecular mechanisms controlling the mode of cell division and cell cycle parameters of progenitor cells. Similarly, the fate of postmitotic cells can be examined using cell-specific fluorescent reporter proteins or post-imaging immunocytochemistry. More importantly, all these features can be analyzed at the single-cell level, allowing the identification of progenitors committed to the generation of specific cell types. Manipulation of gene expression can also be performed using viral-mediated transfection, allowing the study of cell-autonomous and non-cell-autonomous phenomena. Finally, the use of fusion fluorescent proteins allows the study of symmetric and asymmetric distribution of selected proteins during division and the correlation with daughter cells fate. Here, we describe the time-lapse video-microscopy method to image primary cerebral cortex murine cells for up to several days and analyze the mode of cell division, cell cycle length and fate of newly generated cells. We also describe a simple method to transfect progenitor cells, which can be applied to manipulate genes of interest or simply label cells with reporter proteins.

Live imaging is a powerful tool to study cellular behaviors in real time. Here, we describe a protocol for time-lapse video-microscopy of primary cerebral cortex cells that allows a detailed examination of the phases enacted during the lineage progression from primary neural stem cells to differentiated neurons and glia.

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or ...

Exploring Deep Space - Uncovering the Anatomy of Periventricular Structures to Reveal the Lateral Ventricles of the Human Brain

Anatomy students are typically provided with two-dimensional (2D) sections and images when studying cerebral ventricular anatomy and students find this challenging. Because the ventricles are negative spaces located deep within the brain, the only way to understand their anatomy is by appreciating their boundaries formed by related structures. Looking at a 2D representation of these spaces, in any of the cardinal planes, will not enable visualisation of all of the structures that form the boundaries of the ventricles. Thus, using 2D sections alone requires students to compute their own mental image of the 3D ventricular spaces. The aim of this study was to develop a reproducible method for dissecting the human brain to create an educational resource to enhance student understanding of the intricate relationships between the ventricles and periventricular structures. To achieve this, we created a video resource that features a step-by-step guide using a fiber dissection method to reveal the lateral and third ventricles together with the closely related limbic system and basal ganglia structures. One of the advantages of this method is that it enables delineation of the white matter tracts that are difficult to distinguish using other dissection techniques. This video is accompanied by a written protocol that provides a systematic description of the process to aid in the reproduction of the brain dissection. This package offers a valuable anatomy teaching resource for educators and students alike. By following these instructions educators can create teaching resources and students can be guided to produce their own brain dissection as a hands-on practical activity. We recommend that this video guide be incorporated into neuroanatomy teaching to enhance student understanding of the morphology and clinical relevance of the ventricles.

This paper demonstrates the effective use of a fiber dissection method to reveal the superficial white matter tracts and periventricular structures of the human brain, in three-dimensional space, to aid student comprehension of ventricular morphology.

Anatomy students are typically provided with two-dimensional (2D) sections and images when studying cerebral ventricular anatomy and students find this ...

Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical properties, synaptic connections, and in vivo functions, but their basic functional and anatomical organization from cellular to network scale is poorly understood. Here we describe a method for the three-dimensional imaging of fluorescently-labeled neurons across large areas of the brain for the investigation of the cortical cellular organization. Specific types of neurons are labeled by the injection of fluorescent retrograde neuronal tracers or expression of fluorescent proteins in transgenic mice. Block brain samples, e.g., a hemisphere, are prepared after fixation, made transparent with tissue clearing methods, and subjected to fluorescent immunolabeling of the specific cell types. Large areas are scanned using confocal or two-photon microscopes equipped with large working distance objectives and motorized stages. This method can resolve the periodic organization of the cell type-specific microcolumn functional modules in the mouse neocortex. The procedure can be useful for the study of three-dimensional cellular architecture in the diverse brain areas and other complex tissues.

Here we describe a procedure for tissue clearing, fluorescent labeling, and large-scale imaging of mouse brain tissue which, thereby, enables visualization of the three-dimensional organization of cell types in the neocortex.