This work was supported by Humboldt Universit&#228;t zu Berlin, the Bernstein Center for Computational Neuroscience Berlin, the German Center for Neurodegenerative Diseases (DZNE), the German Federal Ministry of Education and Research (BMBF, F&#246;rderkennzeichen 01GQ1001A), NeuroCure, and the Gottfried Wilhelm Leibniz prize of the DFG. We thank Shimpei Ishiyama for excellent graphic design and Juliane Diederichs for excellent technical assistance.

All experimental procedures were performed according to the German guidelines on animal welfare under the supervision of local ethics committees (LaGeSo). Human and bat brain data were derived from Naumann et al.5 The following procedure is performed on a male adult Wistar rat (strain: RJHan:WI).
1. Perfusion and Brain Extraction
NOTE: In order to obtain a homogenously fixed and blood-free brain, transcardial perfusion of the animal is highly encouraged, as residual blood increases unspecific background signal during staining. Nevertheless, it is also possible to obtain flattened sections from un-perfused specimen and stain them. The ease of handling the specimen varies with the concentration of fixative used. Too little fixation increases the risk of handling damage to the brain during flattening and cutting, while too high concentrations lower the flexibility for flattening and the quality of the staining signal.
<ol>
	<li>Transcardially perfuse the animal, using a 23G needle. For a more detailed guide, see Gage et al.9
	<ol>
		<li>Use phosphate buffer saline10 (PBS; 0.02 M; pH 7.4) to flush out blood from the brain and body of the animal. Continue until infusion liquid appears clear, with at least 150 mL PBS being infused through the vascular system. 
		NOTE: Best results are achieved at a pressure similar to the range of the physiological blood pressure of the animal being perfused (e.g., rat: 120-130 mmHg). Optionally, add heparin (10 U/mL) to the solution to avoid blood coagulation11.</li>
		<li>To fix the brain, transcardially infuse a minimum of 100 mL paraformaldehyde12 (PFA, 4%; in 0.1 M phosphate buffer (PB)13) until the neck of the animal is stiff. 
		CAUTION: PFA is a potential carcinogen. 
		NOTE: Best results for histochemical activity staining, such as for cytochrome oxidase, are achieved, when using 2% PFA. For other staining, such as immunohistochemistry, 4% PFA is preferred.</li>
	</ol>
	</li>
	<li>Extract the brain from the skull. 
	NOTE: See Gage et al.9 for details.
	<ol>
		<li>Isolate the head using large scissors or bone shears.</li>
		<li>Using fine scissors, cut the skin along the midline from neck to nose. Pull apart the skin to expose the skull. Remove the neck and temporal muscles using fine scissors.</li>
		<li>Make a midline cut through the interparietal bone starting from the foramen magnum up to the parietal bone.</li>
		<li>Use Rongeurs to peel off the interparietal bone. Slide fine scissors along the sagittal suture and cut from the posterior end of the parietal to the anterior end of the frontal bones.</li>
		<li>Use Rongeurs to peel off the parietal and frontal bones. Alternatively, lift the bones away using forceps. Use a spoon to cut ventral nerve cords and remove the brain. Transfer the brain to PB (0.1 M; pH 7.4).</li>
	</ol>
	</li>
	<li>Fix un-perfused brains by immersion in 4% PFA for 1-3 h at 4 &#176;C (depending on the brain size and probe: Shrew: ~ 1 h, Human: ~ 3 h). 
	NOTE: For large and gyrencephalic brains like humans, isolate and cut out the area of interest to reduce fixation time.</li>
	<li>To obtain larger cortical areas in a section, flatten the brain prior to further post-fixation; proceed to step 2. However, to obtain sections of harder to reach areas like the medial entorhinal cortex, post-fix the brain in 4% PFA for 24 h before proceeding to step 2.</li>
</ol>
2. Brain Dissection and Flattening
<ol>
	<li>Separate the brain hemispheres by using a razor blade (lissencephalic brains) or scalpel (gyrencephalic brains). 
	NOTE: If the brain has not been post-fixed, it is susceptible to damage by handling.</li>
	<li>Optional step for aiding subsequent section registration and shrinkage estimation:
	<ol>
		<li>Puncture the cortex in an area of non-interest, with a 35G needle normal to the cortical surface. Repeat the step two times at defined distances along the cortical sheet. To determine the distance along the cortical sheet, use a pre-cut thread at a fixed length (e.g., 5 mm) and lay it along the cortical surface to determine points of puncture.</li>
	</ol>
	</li>
	<li>Lissencephalic brains: Remove subcortical structures by using a dissecting spatula. Critical: Keep the brain moist with PB (0.1 M; pH 7.4) throughout the whole procedure.
	<ol>
		<li>Hold the brain by the cerebellum and gently insert the spatula to open a dissection plane in the corpus callosum. The round tip of spatula should point away from the cortex.</li>
		<li>Slide the spatula further and pull gently until the spatula is between the thalamus and cortex. Separate subcortical structures with scratching motions.</li>
		<li>Inspect the hemisphere for even thickness. 
		NOTE: Any uneven regions might result in a trans-layer gradient during sectioning. For tangential sectioning, use a scalpel to make a clean cut through the brain, parallel to the pial surface of the region from which tangential sections are desired.</li>
		<li>Optional steps to improve flattening quality:
		<ol>
			<li>Make a clean cut through the striatum, nucleus accumbens, and orbitofrontal cortex, since they increase the thickness of the cortex in the lateral region. Also, add a relieving cut at the base of the inner region of prefrontal cortex, which allows unfolding of medial portions of the cortex.</li>
		</ol>
		</li>
		<li>Place the hemisphere (cortex facing down) on a glass slide. Proceed to step 2.6.</li>
	</ol>
	</li>
	<li>Gyrencephalic brains: Remove white matter to unfold gyri (for detailed protocols, see: Sincich et al.14; Tootell and Silverman15). Critical: Use PB (0.1 M; pH 7.4) to keep the brain moist at all time.
	<ol>
		<li>Put the area of interest on a damp filter paper in a large Petri dish, with the cortex facing up.</li>
		<li>Remove the arachnoid membrane and pia using two forceps.</li>
		<li>Use a damp cotton swab and insert gently in each sulcus to detach adhesions between gyri.</li>
		<li>Turn the brain around by using another damp filter paper.</li>
		<li>Use two curved micro spatulas to unfold single gyri. Use the convex side of the spatulas to tease apart the white matter until reaching close to the concave end of the gyrus. Use a damp cotton swab and proceed with swirling motions to reach the concave end of the gyrus.</li>
		<li>Tease apart single gyri. If necessary, add small relieving cuts if the tension is too high.</li>
		<li>Flatten the unfolded brain in the Petri dish or a similar container.</li>
	</ol>
	</li>
	<li>Optionally, place larger brains on a filter paper covered sponge, to ensure that regions do not dry out.</li>
	<li>Place two rolled pieces of clay on both sides. Critical: As the thickness of the clay defines how much the hemisphere can be flattened, ensure that the clay is 10-20% thinner than the un-flattened cortex.</li>
	<li>Gently press a second glass slide/small Petri dish on the cortex until the hemisphere is fully flat. To obtain the best results for a desired region, place the glass slide first on the respective area. To obtain the best results for lissencephalic brains, place the glass slide tangentially at the lateral portion first and apply concentrated pressure on this region.</li>
	<li>Put a weight (e.g., a ceramic watch glass) on the glass slides/Petri dish and flatten the hemisphere for 3-5 h at 4 &#176;C in PB.</li>
	<li>For post-fixation, release the pressure and remove the glass slides. Put the flattened hemisphere in PFA (free floating) for 24 h on a shaker at 4 &#176;C. 
	NOTE: Best results for histochemical activity stainings are achieved when using 1% PFA. For other stainings, such as immunohistochemistry, for un-perfused brains, 2% PFA can be used.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56992/56992fig1.jpg" /> 
Figure 1: Schematic representation of workflow for flattening of a rat cortical hemisphere and visualization of modules in somatosensory cortex. Subsequent to the transcardial perfusion, the brain of a mouse was dissected (A). Subcortical structures were removed and cortex was flattened between two glass slides in phosphate buffer (B). Flattened hemisphere (C) was post-fixed, tangentially sectioned, and stained for cytochrome oxidase activity (D). Scale bars = 1 cm. R: Rostral, C: Caudal, L: Lateral, M: Medial. Figure adapted from Lauer et al.23 <a href="//cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56992/56992fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Cutting Tangential Sections
NOTE: Depending on the requirements of the staining protocols, the cutting procedure and thickness can be adapted. A vibratome was used to cut the hemispheres for further histochemical processing (step 3.2) at 80-150 &#181;m. However, for immunohistochemical processing, thinner sections are desired and a freezing microtome was used for sectioning (step 3.3) at 10-60 &#181;m. See Video 1.
<ol>
	<li>Wash the flattened hemisphere in PB (0.1M; pH 7.4) for 15 min.</li>
	<li>Cut the hemisphere on a vibratome.
	<ol>
		<li>Place the hemisphere tangentially on the slicing holder. Optionally, press again gently with a glass slide before fixing it in position (e.g., with superglue). 
		NOTE: Minimize the amount of glue used, because cutting artifacts could result due to the small thickness of flattened hemispheres.</li>
		<li>Cut the hemisphere from the thicker and more stable end towards the thinner end (posterior to anterior, here) at the required thickness. Proceed to step 3.4. 
		NOTE: If low fixation concentrations were used, cut it at a slow speed and a high amplitude.</li>
	</ol>
	</li>
	<li>Cut the hemisphere on a freezing microtome.
	<ol>
		<li>Cryoprotect the brain by immersing it in a 30% sucrose solution (in PB) until it sinks. 
		NOTE: Depending on the size of the tissue being cryoprotected, sinking in the solution can range from a few hours to a few days. For larger brains alternative cryoprotection methods might be considered (see Rosene et al.16).</li>
		<li>Form an ice base on the freezing microtome to mount the brain. Construct the ice base by freezing PB on the block face of the freezing microtome. Critical: Ensure that the surface of the ice base is parallel to the microtome blade. To do this, cut the empty ice base using the microtome blade to exactly align the ice base parallel to the cutting surface.</li>
		<li>Embed the brain in freezing medium and mount it to the block face of the microtome with the area of interest parallel to the block face. Face the pial surface of the brain towards the block face of the microtome. Critical: Adjust the freezing temperature depending on sample size; higher temperatures ensure better section integrity while sectioning, but larger sections require lower temperatures to freeze sample uniformly.</li>
		<li>Cut the brain tangentially at the required thickness (slower and uniform cutting speeds result in best section quality).</li>
	</ol>
	</li>
	<li>Wash the sections in PB for 15 min on a shaker.</li>
</ol>
<img alt="Video Snapshot" src="/files/ftp_upload/56992/56992video1.jpg" /> 
Video 1: Schematic video of tangential sectioning from a rat medial entorhinal cortex and layout of parasubicular and entorhinal modules. The medial entorhinal cortex of a rodent brain is situated at the posterior end of the cortex and is tilted towards the medial and ventral side. Tangential sections are obtained by orienting a knife along this angle. Consequently, appropriate cell-type specific staining reveals modular structures in the medial entorhinal cortex and adjoining parasubiculum. Video adapted from Ray and Brecht8. <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56992/Video_1_Tangential_Procedure_MEC.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
4. Visualization of Cortical Modules Using Cytochrome Oxidase Staining
NOTE: Different staining protocols have been developed for histochemical detection of cytochrome oxidase activity, e.g., first by Wong-Riley17 and later modified by Divac et al.18 This protocol is based on the one by Divac et al.18, since the use of nickel-ammonium sulfate (NiAS) results in a higher contrast and better defined modules in stained cortical areas.
<ol>
	<li>Prepare the cytochrome oxidase staining solution (see Divac et al.18). For 10 mL of solution, add: 10 mL HEPES buffer (0.1 M, pH 7.4), 400 mg sucrose, 12.5 mg NiAS, 2 mg cytochrome C, 6 mg diaminobenzidine (DAB). 
	CAUTION: DAB and NiAS are carcinogenic. 
	NOTE: Add DAB just prior to incubation of sections.</li>
	<li>Wash the sections in HEPES buffer (0.1 M, pH 7.4) on a shaker for 15 min.</li>
	<li>Incubate the sections in the staining solution at room temperature on a shaker. 
	NOTE: Observe the speed of the staining. If there is no visible reaction, change to incubation at 37 &#176;C. Depending on the amount of fixation, staining can be observed after 10 min or in several hours.</li>
	<li>Stop the reaction by adding 4% PFA; this prevents unwanted re-dying and increase of background signal.</li>
	<li>Wash the sections three times using HEPES buffer for 10 min.</li>
	<li>Mount and dry the sections on glass slides.</li>
	<li>Dehydrate the sections with an increasing alcohol row:
	<ol>
		<li>Wash the slides in 60% ethanol for 1 min. Wash the slides in 80% ethanol for 1 min. Wash the slides in 96% ethanol for 2 min. Wash the slides in 100% ethanol for 3 min. Wash the slides in isopropanol for 5 min. Wash the slides in xylene for 5 min.</li>
	</ol>
	</li>
	<li>Immediately add a quick hardening-mounting medium, and add a coverslip. Critical: Do not use water-based mounting mediums, as NiAS will be washed out and degrade the staining.</li>
	<li>Keep the sections at 4 &#176;C for long term storage.</li>
</ol>
5. Visualization of Cortical Modules Using Immunohistochemical Staining
NOTE: Multiple protocols are available for immunohistochemistry, optimized for the specimen and the type of probe. Adaptations can be made as required, by varying concentrations of antibodies, permeabilizing agents, and incubation times. The following protocol leads to good results for detecting a large range of antibodies and visualization by fluorescent probes.
<ol>
	<li>Wash the sections in PBS (0.1 M; pH 7.4) for 15 min.</li>
	<li>Optional: Perform antigen retrieval to unmask an epitope using a water bath (based on Jiao et al.19; for alternative methods, see Pileri et al.20).
	<ol>
		<li>Preheat a water bath to 80 &#176;C. Prepare tri-sodium citrate buffer19 (pH 8.0) and preheat it in the water bath to 80 &#176;C.</li>
		<li>Transfer the sections to tri-sodium citrate buffer. Incubate the sections for 30 min at 80 &#176;C</li>
		<li>Cool the sections down to room temperature</li>
		<li>Wash the sections in PBS for 15 min.</li>
	</ol>
	</li>
	<li>Wash the sections two times in PBS-X (0.5% Triton-X, in 0.1 M PBS) for 15 min.</li>
	<li>Block unspecific epitopes by incubating the sections in a solution of Bovine Serum Albumin (BSA; 2.5%) and Triton-X (0.75%) in PBS for 2 h.</li>
	<li>Incubate the sections in primary antibody, e.g., Calbindin- D28k (1:5,000, 1% BSA in PBS-X), for 2-3 days on a shaker at 4 &#176;C. 
	NOTE: Optimal dilution for the primary antibody depends on the specific antibody used. Refer to manufacturer&#39;s information to obtain dilution criteria for a specific antibody. Multiple antibodies can be used together but must be raised against different species.</li>
	<li>Wash the sections three times in PBS for 15 min each.</li>
	<li>Incubate the sections overnight in secondary antibody (1:200, 1% BSA in PBS) at 4 &#176;C on a shaker. 
	NOTE: Multiple secondary antibodies can be used at different spectra if multiple primary antibodies were used.</li>
	<li>Wash off the secondary antibody by washing the sections three times in PBS for 10 min each.</li>
	<li>Mount and dry the sections on a glass slide for microscopy. 
	NOTE: The sections should still be moist prior to mounting the coverslip.</li>
	<li>Apply mounting medium suited for fluorescence dyes on the tissue sections and apply a coverslip.</li>
	<li>Dry the sections for 1 h.</li>
	<li>For long term storage, seal the coverslip using nail polish and keep in the dark at 4 &#176;C.</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thalamic+inputs+to+cytochrome+oxidase-rich+regions+in+monkey+visual+cortex.">Thalamic inputs to cytochrome oxidase-rich regions in monkey visual cortex.</a> Proc Natl Acad Sci U S A. 79 (19), 6098-6101 (1982).</li><li>Land, P. W., Simons, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytochrome+oxidase+staining+in+the+rat+SmI+barrel+cortex.">Cytochrome oxidase staining in the rat SmI barrel cortex.</a> J Comp Neurol. 238 (2), 225-235 (1985).</li><li>Welker, C., Woolsey, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+of+layer+IV+in+the+somatosensory+neocortex+of+the+rat:+description+and+comparison+with+the+mouse.">Structure of layer IV in the somatosensory neocortex of the rat: description and comparison with the mouse.</a> J Comp Neurol. 158 (4), 437-453 (1974).</li><li>Retzius, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Die+Cajal'schen+zellen+der+grosshirnrinde+beim+menschen+und+bei+s&#228;ugetieren.">Die Cajal'schen zellen der grosshirnrinde beim menschen und bei s&#228;ugetieren.</a> Biol Unters. 5, 1-9 (1893).</li><li>Cajal, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Histologie du Systeme Nerveux de l'Homme et des vert&#233;br&#233;s. , (1911).</li><li>Chapin, J. K., Lin, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+the+body+representation+in+the+SI+cortex+of+anesthetized+and+awake+rats.">Mapping the body representation in the SI cortex of anesthetized and awake rats.</a> J Comp Neurol. 229 (2), 199-213 (1984).</li><li>L&#246;wel, S., Freeman, B., Singer, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Topographic+organization+of+the+orientation+column+system+in+large+flat-mounts+of+the+cat+visual+cortex:+A+2-deoxyglucose+study.">Topographic organization of the orientation column system in large flat-mounts of the cat visual cortex: A 2-deoxyglucose study.</a> J Comp Neurol. 255 (3), 401-415 (1987).</li><li>Tang, 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=Functional+architecture+of+the+rat+parasubiculum.">Functional architecture of the rat parasubiculum.</a> J Neurosci. 36 (7), 2289-2301 (2016).</li><li>Snyder, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Map projections--A working manual (Vol. 1395). , (1987).</li><li>Chung, K., Deisseroth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CLARITY+for+mapping+the+nervous+system.">CLARITY for mapping the nervous system.</a> Nat Methods. 10 (6), 508-513 (2013).</li><li>Renier, N., Wu, Z., Simon, D. J., Yang, J., Ariel, P., Tessier-Lavigne, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=iDISCO:+a+simple,+rapid+method+to+immunolabel+large+tissue+samples+for+volume+imaging.">iDISCO: a simple, rapid method to immunolabel large tissue samples for volume imaging.</a> Cell. 159 (4), 896-910 (2014).</li></ol>

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Modularity in the cerebral cortex has been identified using a variety of techniques. The earliest studies typically identified cortical modules by either visualizing cell dense regions, or an absence of fibers1. Subsequent methods have utilized the presence of dendritic bundles24, afferents from a particular region25, or enrichment of neurotransmitters26. Here we demonstrate two techniques, (i) cytochrome oxidase histochemistr...

Some of the most significant changes in the brain structure across phylogeny can be observed in the cerebral cortex. Despite significant differences, the cortex of animals follows a common pattern and can be broadly divided in two distinct ways, by layers and areas1. Cortical layers lie parallel to the surface of the brain and vary in number from 3 layers in reptilian cortices2 to 6 layers in mammalian cortices1. Cortical areas on the other hand are distinct regions of the cortex which largely correspond to distinct functionalities, e.g., the somatosensory cortex is involved in the sensation of touch or the visual cortex in processing visual inputs. These cortical areas can often be subdivided into patches or modules3, which are regularly repeating anatomical structures, essentially found parallel to the pial surface of the brain. Cortical modules may be confined to a particular layer4, or extend across several layers5.
Standard sectioning methods of the brain involve sections normal to the surface of the brain, like coronal or sagittal. While these methods can be used to visualize cortical modules, a multitude of interesting features can be revealed when the cortical modules are visualized tangentially, in a plane parallel to the surface of the brain. For instance, somatosensory modules in the rodent brain representing whiskers, appear as barrels when visualized normal to the brain surface, and thus the regions derive the name barrel cortex. However, on visualizing the barrels in a tangential orientation, they reveal a whisker-map, with the barrels being laid out in a topographic orientation mirroring the exact layout of the whiskers on the external body surface. In certain cases, modular arrangement has even escaped detection for considerable periods, when visualized in a non-tangential manner. The medial entorhinal cortex, is known for the presence of grid cells, neurons which fire in a regular hexagonal pattern when an animal is traversing an environment. Even though it is a heavily investigated area, until recently, the presence of patches or modules of cells in the medial entorhinal cortex, which are physically laid out in a hexagonal pattern6, had escaped detection. The presence and arrangement of these modules, in the rat brain, was facilitated by making tangential sections of the medial entorhinal cortex and investigating the cytoarchitecture in a layer-wise manner.
Subsequent to sectioning, the particular aspect of visualization of cortical modules can also be realized in multiple ways. Classically, studies have delineated modules based on cell density or fiber layout1. Another popular approach is the use of cytochrome oxidase histochemistry, which reveals areas of higher activity8. Newer approaches include looking at genetically determined cell types, distinguished on the basis of their protein expression profiles6,8.
In this study, we highlight methods to isolate the cortex from mammalian brains, obtain flattened tangential sections, and visualize cortical modules based on cytochrome oxidase histochemistry and immunohistochemistry of cell-type specific proteins.

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			<td>Carl Roth</td>
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			<td>Immunohistochemistry</td>
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			<td>Calbindin D-28k from recombinant rat calbindin D-28k, Rabbit polyclonal</td>
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			<td>Eukitt (histochemistry)</td>
			<td>Sigma-Aldrich</td>
			<td>03989</td>
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			<td>Tissue freezing medium</td>
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			<td>Carl Roth</td>
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			<td>Carl Roth</td>
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			<td>Devices/tools</td>
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			<td>Thermo Scientific</td>
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			<td>Leica Biosystems</td>
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			<td>Dumont #55 Forceps - Inox</td>
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			<td>11255-20</td>
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			<td>Dumont #5 Forceps - Inox Biology Tip</td>
			<td>Fine Science Tools</td>
			<td>11252-30</td>
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			<td>Fine Science Tools</td>
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			<td>Fine Science Tools</td>
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			<td>Fine Science Tools</td>
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			<td>Surgical Scissors - Sharp-Blunt</td>
			<td>Fine Science Tools</td>
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			<td>Fine Iris Scissors</td>
			<td>Fine Science Tools</td>
			<td>14094-11</td>
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visualization of cortical modules in flattened mammalian cortices

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and can be delineated by certain histochemical and immunohistochemical methods. In this study, we highlight a method to isolate the cortex from mammalian brains and flatten them to obtain sections parallel to the cortical sheet. We further highlight selected histochemical and immunohistochemical methods to process these flattened tangential sections to visualize cortical modules. In the somatosensory cortex of various mammals, we perform cytochrome oxidase histochemistry to reveal body maps or cortical modules representing different parts of the body of the animal. In the medial entorhinal cortex, an area where grid cells are generated, we utilize immunohistochemical methods to highlight modules of genetically determined neurons which are arranged in a grid-pattern in the cortical sheet across several species. Overall, we provide a framework to isolate and prepare layer-wise flattened cortical sections, and visualize cortical modules using histochemical and immunohistochemical methods in a wide variety of mammalian brains.

We obtained flattened cortical sections of the somatosensory cortex in a variety of brains, and processed them for cytochrome oxidase histochemistry to visualize the somatotopic modules representing different body parts. This comparative approach allows studying the evolutionary forces that shape cortex, e.g., showing highly conserved representation of mystacial vibrissae in rodents and lagomorpha as barrels21 (Figure 2). In c...

Watch this Scientific Journal Video about Visualization of Cortical Modules in Flattened Mammalian Cortices at JoVE.com

Visualization of Cortical Modules in Flattened Mammalian Cortices

This article describes a detailed methodology to obtain flattened tangential sections from mammalian cortices and visualize cortical modules using histochemical and immunohistochemical methods.

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and ...

visualization-of-cortical-modules-in-flattened-mammalian-cortices

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Neuroscience

12.7K Views.  Humboldt University of Berlin. This article describes a detailed methodology to obtain flattened tangential sections from mammalian cortices and visualize cortical modules using histochemical and immunohistochemical methods.

Video: Visualization of Cortical Modules in Flattened Mammalian Cortices

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Automated Interactive Video Playback for Studies of Animal Communication

Video playback is a widely-used technique for the controlled manipulation and presentation of visual signals in animal communication. In particular, parameter-based computer animation offers the opportunity to independently manipulate any number of behavioral, morphological, or spectral characteristics in the context of realistic, moving images of animals on screen. A major limitation of conventional playback, however, is that the visual stimulus lacks the ability to interact with the live animal. Borrowing from video-game technology, we have created an automated, interactive system for video playback that controls animations in response to real-time signals from a video tracking system. We demonstrated this method by conducting mate-choice trials on female swordtail fish, Xiphophorus birchmanni. Females were given a simultaneous choice between a courting male conspecific and a courting male heterospecific (X. malinche) on opposite sides of an aquarium. The virtual male stimulus was programmed to track the horizontal position of the female, as courting males do in the wild. Mate-choice trials on wild-caught X. birchmanni females were used to validate the prototype's ability to effectively generate a realistic visual stimulus.

Video playback is a widely used technique in animal behavior. We created and evaluated a program that applies rules-based, interactive playback of 3-D computer animations in response to real-time, automated data on subject behavior.

Video playback is a widely-used technique for the controlled manipulation and presentation of visual signals in animal communication. In particular, ...

Functional Calcium Imaging in Developing Cortical Networks

A hallmark pattern of activity in developing nervous systems is spontaneous, synchronized network activity. Synchronized activity has been observed in intact spinal cord, brainstem, retina, cortex and dissociated neuronal culture preparations. During periods of spontaneous activity, neurons depolarize to fire single or bursts of action potentials, activating many ion channels. Depolarization activates voltage-gated calcium channels on dendrites and spines that mediate calcium influx. Highly synchronized electrical activity has been measured from local neuronal networks using field electrodes. This technique enables high temporal sampling rates but lower spatial resolution due to integrated read-out of multiple neurons at one electrode. Single cell resolution of neuronal activity is possible using patch-clamp electrophysiology on single neurons to measure firing activity. However, the ability to measure from a network is limited to the number of neurons patched simultaneously, and typically is only one or two neurons. The use of calcium-dependent fluorescent indicator dyes has enabled the measurement of synchronized activity across a network of cells. This technique gives both high spatial resolution and sufficient temporal sampling to record spontaneous activity of the developing network.
A key feature of newly-forming cortical and hippocampal networks during pre- and early postnatal development is spontaneous, synchronized neuronal activity (Katz &amp; Shatz, 1996; Khaziphov &amp; Luhmann, 2006). This correlated network activity is believed to be essential for the generation of functional circuits in the developing nervous system (Spitzer, 2006). In both primate and rodent brain, early electrical and calcium network waves are observed pre- and postnatally in vivo and in vitro (Adelsberger et al., 2005; Garaschuk et al., 2000; Lamblin et al., 1999). These early activity patterns, which are known to control several developmental processes including neuronal differentiation, synaptogenesis and plasticity (Rakic &amp; Komuro, 1995; Spitzer et al., 2004) are of critical importance for the correct development and maturation of the cortical circuitry.
In this JoVE video, we demonstrate the methods used to image spontaneous activity in developing cortical networks. Calcium-sensitive indicators, such as Fura 2-AM ester diffuse across the cell membrane where intracellular esterase activity cleaves the AM esters to leave the cell-impermeant form of indicator dye. The impermeant form of indicator has carboxylic acid groups which are able to then detect and bind calcium ions intracellularly.. The fluorescence of the calcium-sensitive dye is transiently altered upon binding to calcium. Single or multi-photon imaging techniques are used to measure the change in photons being emitted from the dye, and thus indicate an alteration in intracellular calcium. Furthermore, these calcium-dependent indicators can be combined with other fluorescent markers to investigate cell types within the active network.

Spontaneous activity of developing neuronal networks can be measured using AM-ester forms of calcium-sensitive indicator dyes. Changes in intracellular calcium, indicating neuronal activation, are detected as transient changes in indicator fluorescence with one- or two-photon imaging. This protocol can be adapted for a range of developmentally-dependent neuronal networks in vitro.

A hallmark pattern of activity in developing nervous systems is spontaneous, synchronized network activity. Synchronized activity has been observed in ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Isolation and Culture of Mouse Cortical Astrocytes

Astrocytes are an abundant cell type in the mammalian brain, yet much remains to be learned about their molecular and functional characteristics. In vitro astrocyte cell culture systems can be used to study the biological functions of these glial cells in detail. This video protocol shows how to obtain pure astrocytes by isolation and culture of mixed cortical cells of mouse pups. The method is based on the absence of viable neurons and the separation of astrocytes, oligodendrocytes and microglia, the three main glial cell populations of the central nervous system, in culture. Representative images during the first days of culture demonstrate the presence of a mixed cell population and indicate the timepoint, when astrocytes become confluent and should be separated from microglia and oligodendrocytes. Moreover, we demonstrate purity and astrocytic morphology of cultured astrocytes using immunocytochemical stainings for well established and newly described astrocyte markers. This culture system can be easily used to obtain pure mouse astrocytes and astrocyte-conditioned medium for studying various aspects of astrocyte biology.

Astrocytes have been recognized to be versatile cells participating in fundamental biological processes that are essential for normal brain development and function, and central nervous system repair. Here we present a rapid procedure to obtain pure mouse astrocyte cultures to study the biology of this major class of central nervous system cells.

Astrocytes  are an abundant cell type in the mammalian brain, yet much remains to be  learned about their molecular and functional characteristics. In ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Subtype-selective Electroporation of Cortical Interneurons

The study of central nervous system (CNS) maturation relies on genetic targeting of neuronal populations. However, the task of restricting the expression of genes of interest to specific neuronal subtypes has proven remarkably challenging due to the relative scarcity of specific promoter elements. GABAergic interneurons constitute a neuronal population with extensive genetic and morphological diversity. Indeed, more than 11 different subtypes of GABAergic interneurons have been characterized in the mouse cortex1. Here we present an adapted protocol for selective targeting of GABAergic populations. We achieved subtype selective targeting of GABAergic interneurons by using the enhancer element of the homeobox transcription factors Dlx5 and Dlx6, homologues of the Drosophila distal-less (Dll) gene2,3, to drive the expression of specific genes through in utero electroporation.

This procedure shows how to target interneurons in the developing mouse forebrain by means of in utero electroporation. This technique was particularly efficient to achieve selective gene expression in interneuron subtypes destined to the superficial layers of the cortex.

The study of central nervous system (CNS) maturation relies on genetic targeting of neuronal populations. However, the task of restricting the expression ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) axons form branches and synapses in specific layers of the cerebral cortex. Despite the obvious spatial correlation between axon branching and synapse formation, the causal relationship between them is poorly understood. To address this issue, we recently developed a method for simultaneous imaging of branching and synapse formation of individual TC axons in organotypic cocultures.
This protocol describes a method which consists of a combination of an organotypic coculture and electroporation. Organotypic cocultures of the thalamus and cerebral cortex facilitate gene manipulation and observation of axonal processes, preserving characteristic structures such as laminar configuration. Two distinct plasmids encoding DsRed and EGFP-tagged synaptophysin (SYP-EGFP) were co-transfected into a small number of thalamic neurons by an electroporation technique. This method allowed us to visualize individual axonal morphologies of TC neurons and their presynaptic sites simultaneously. The method also enabled long-term observation which revealed the causal relationship between axon branching and synapse formation.

This protocol describes a method for simultaneous imaging of thalamocortical axon branching and synapse formation in organotypic cocultures of the thalamus and cerebral cortex. Individual thalamocortical axons and their presynaptic terminals are visualized by a single cell electroporation technique with DsRed and GFP-tagged synaptophysin.

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) ...