We thank Irmgard Weiss for excellent technical assistance. The authors acknowledge support by Hermes Pofantis and Andoniya Petkova. The authors also thank the European Neuroscience Institute for the usage of the LSM800 and technical assistance, especially by Dr. Nils Halbsgut. This work was funded by the University Medical Center G&#246;ttingen. JSV acknowledges support by the Center for Nanoscale Microscopy and Molecular Physiology of the Brain (CNMPB).

This protocol does not involve experiments on live animals. Experiments involving euthanizing of animals to obtain brain samples were approved by the local animal protection authorities (Tierschutzkommission der Universit&#228;tsmedizin G&#246;ttingen) under the approval number T 10/30.
NOTE: For this protocol, 3 adult male C57BL/6 mice were used.
1. Sample Preparation
<ol>
	<li>Prepare fixative and 0.1 M phosphate buffer (PB; see Table 1).</li>
	<li>Fix the animal by transcardial perfusion as described in Gage et al.28. First wash out the blood with 0.9% NaCl-solution, then perfuse with 30 mL of 4% paraformaldehyde (PFA).</li>
	<li>Open the skull with scissors and carefully isolate the brain using a spoon with blunt edges to avoid damaging the tissue.</li>
	<li>Fill a 50 mL reaction tube with fixative and postfix the brain in 4% PFA at 4 &#176;C overnight.</li>
	<li>Remove the fixative and wash the brain in 50 mL of 0.1 M PB on a shaker for 30 min.</li>
	<li>After washing, incubate the brain in a 50 mL reaction tube in 30% sucrose in 0.1 M PB for 48 h or until it sinks in the tube at 4 &#176;C for cryoprotection.</li>
	<li>Trim the cryoprotected brain with a sharp blade, place it in a cryomold, and embed it with optimal cutting temperature (OCT) compound. Avoid bubbles. Orient the brain and freeze the cryomold in the -80 &#176;C freezer.</li>
	<li>Mount the frozen tissue for sectioning. Equilibrate the tissue to the cryomicrotome temperature for at least 15 min before sectioning.</li>
	<li>Section the brain into 25 &#181;m thick coronal slices. Touch the OCT carefully with a glass hook without touching the brain tissue. Collect 3 adjacent slices per well in a 24 well plate and store them in 0.1 M PB at 4 &#176;C until staining. 
	NOTE: The protocol can be paused here for up to two weeks. Longer storage times can interfere with the tissue quality and thus influence the outcome of the experiment.</li>
</ol>
2. Immunofluorescence
<ol>
	<li>Prepare solutions including the blocking buffer, antibody buffer, washing buffer 1, and washing buffer 2 (see Table 1).</li>
	<li>Rinse slices once with PB to remove excess OCT.
	<ol>
		<li>Remove the solution with a plastic pipette without sucking in the brain slices. Add 250 &#181;L of fresh PB with a 1000 &#181;L pipette. 
		CAUTION: Slices should not dry out, so remove and add fluids well by well.</li>
	</ol>
	</li>
	<li>Remove the PB with a plastic pipette and add 250 &#181;L of blocking buffer per well with a 1000 &#181;L pipette. Incubate for 3 h at room temperature (RT) on the shaker.</li>
	<li>During the incubation time, dilute the primary antibodies in antibody buffer in a reaction tube. Use 250 &#181;L antibody buffer per well and add the appropriate amount of antibody (see Table 2) by pipetting it directly into the solution using a 2 &#181;L pipette. Mix the solution by gently pipetting up and down several times. Vortex shortly afterwards to ensure proper mixing. 
	NOTE: To determine the background fluorescence, stainings should also be performed without adding the primary antibody. For that, incubate the slice in antibody solution without primary antibodies according to the protocol.</li>
	<li>After the incubation time, remove the blocking buffer with a plastic pipette and add 250 &#181;L of antibody solution containing primary antibodies per well. Incubate slices with primary antibody overnight at 4 &#176;C on a shaker.</li>
	<li>Next day, wash the slices with washing buffer 1 3x for 10 min at RT on a shaker.
	<ol>
		<li>Remove the medium with a plastic pipette and add 300 &#181;L of washing buffer 1 per well. Incubate at RT for 10 min. Repeat 3 times.</li>
	</ol>
	</li>
	<li>During the washing steps, dilute the fluorophore-coupled secondary antibodies in antibody buffer in a reaction tube. Use 250 &#181;L antibody buffer per well and add the appropriate amount of antibody (see Table 2) by pipetting it directly into the solution using a 2 &#181;L pipette. Mix the solution by gently pipetting up and down several times. Vortex shortly afterwards to ensure proper mixing. 
	CAUTION: Because the antibodies are light-sensitive, all steps from this point on need to be performed in the dark.</li>
	<li>After the washing steps, remove the washing buffer with a plastic pipette and add 250 &#181;L of antibody solution containing secondary antibodies per well. Incubate the slices with secondary antibody for 90 min at RT in the dark.</li>
	<li>Wash the slices with washing buffer 2 3x for 10 min at RT.</li>
	<li>During the washing steps, dilute 4&#8242;,6-diamidino-2-phenylindole (DAPI) in 0.1 M PB in a concentration of 1:2000.</li>
	<li>Remove the washing buffer 2 with a plastic pipette and add 250 &#181;L of DAPI solution per well. Incubate for 5 min at RT on the shaker.</li>
	<li>Remove the DAPI solution with a plastic pipette and add 500 &#181;L of 0.1 M PB per well with a 1000 &#181;L pipette.</li>
	<li>Mount slices on microscope slides.
	<ol>
		<li>Place a microscope slide under the stereoscope. With a fine brush, add three separate drops of 0.1 M PB onto the slide. Place one slice per drop onto the microscope slide.</li>
		<li>Use the fine brush to flatten and orient the slices on the microscope slide.</li>
		<li>When all slices are positioned correctly, remove excess PB with a tissue and dry the slide carefully. 
		CAUTION: Avoid drying the brain slices completely.</li>
		<li>Add 80 &#181;L of embedding medium onto the slide. Carefully lower the coverslip onto the slide, thereby embedding the brain slices.</li>
		<li>Leave the slides to dry in the fume hood for 1-2 h (cover them to avoid light exposure) and store them in a microscope slide box at 4 &#176;C. 
		NOTE: The protocol can be paused here.</li>
	</ol>
	</li>
</ol>
3. Imaging
<ol>
	<li>After the embedding medium is completely hardened, place the microscope slide under the confocal microscope. 
	NOTE: Epifluorescence microscopy combined with deconvolution software should yield similar image quality.</li>
	<li>Adjust the laser settings by increasing or decreasing the laser intensity for every channel so that few pixels are overexposed to ensure maximum distribution of grey values.</li>
	<li>Acquire virtual tissues of the whole brain slice for the different channels.
	<ol>
		<li>In the imaging software (see Table of Materials), select the Tiles option and manually delineate the brain slice with the Tile Region Setup.</li>
		<li>Distribute support points throughout the tile region and adjust the focus for the different support points by pressing Verify Tile Regions/Positions&#8230;.</li>
		<li>Adjust the settings in Acquisition Mode according to the desired resolution and file size of the resulting image and start the scan.</li>
	</ol>
	</li>
	<li>When the scan is finished, use the Stitching function to process the virtual tissue. Export the file as a .tif with the function Image Export.</li>
</ol>
4. Computer-based Analysis
<ol>
	<li>Load all single channels for one image into FIJI29 by clicking File| Open.</li>
	<li>With the Freehand selection tool, delineate one hemisphere in the DAPI-channel. Create a mask of the selection by clicking Edit| Selection| Create mask.</li>
	<li>Determine the mean fluorescence intensity for the single channels (Mover and synaptophysin) by clicking Analyze| Measure. 
	NOTE: Make sure to select the different channels to determine the mean fluorescence intensity values for each channel.</li>
	<li>Copy the mean fluorescence intensity for the single channels into a spreadsheet.</li>
	<li>Determine the mean fluorescence intensity for the single channels in an area of interest by delineating the area also with the Freehand selection tool. Use a mouse brain atlas as reference.</li>
	<li>Repeat steps 4.1-4.5 for all hemispheres and all areas of interest. 
	NOTE: Determine the values for each hemisphere separately in order to later compare the values in an area of interest to that in the hemisphere (see step 5.2).</li>
</ol>
5. Data Handling
<ol>
	<li>In case the background fluorescence is high (see Discussion), a background subtraction might be needed. For that, determine the mean fluorescence intensity for the slice processed without primary antibody against the reference protein (here: synaptophysin) and subtract that value from all values obtained for the brain regions and hemispheres.</li>
	<li>When the mean fluorescence intensities for the single channels for every hemisphere and every area of interest have been determined (see Table 3), calculate the ratio of Mover to synaptophysin by dividing the value for Mover by the value for synaptophysin (yellow in Table 3). Perform this action for every hemisphere and every area of interest separately.</li>
	<li>Divide the ratio obtained for one area of interest by the ratio obtained for the corresponding hemisphere (orange in Table 3) to determine the ratio of the area of interest to the hemisphere.</li>
	<li>To determine the relative Mover abundance, translate the ratio obtained in 5.2 into a percentage by determining its deviation from 1 (red in Table 3). A ratio of 1.25 would therefore give a relative Mover abundance of 25% above average, and a ratio of 0.75 would yield a relative Mover abundance of 25% below average.</li>
</ol>

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The authors have nothing to disclose.

The method presented here aims at quantifying the distribution of a protein of interest relative to the abundance of a marker protein with a known distribution. Immunofluorescence staining can show a high variability of staining intensities between different slices. The quantification approach described here circumvents this problem by determining the ratio of the protein of interest to the average across the hemisphere. Therefore, different staining intensities across slices are cancelled out and allow for a quantitativ...

Communication between neurons happens at specialized contact sites called synapses. Synapses contain a myriad of different proteins that orchestrate synaptic transmission. Some of those proteins show a heterogeneous distribution throughout the nervous system and are not present in every synapse1. One example for such a protein is Munc13, which is involved in the priming process of synaptic vesicles. There are different isoforms of Munc13, which are heterogeneously distributed throughout the brain2, and the presence or absence of specific isoforms can influence short-term synaptic plasticity and synaptic vesicle dynamics3,4,5. Therefore, it is of vital importance to be able to identify the presence of different synaptic proteins across brain areas.
The methods of choice for quantification of synaptic proteins - so far - are mass spectrometry and Western blotting, rather than immunohistochemistry6,7,8,9. In some cases, several methods are used to complement each other to assess both the quantity and the localization of specific proteins (i.e., Wilhelm et al.10). The method we describe here allows for the localization and quantification of proteins of interest without the need of using any biochemical method, simply employing immunofluorescent stainings. Another advantage here is that the quantification can be done over areas much smaller and, therefore, more specific, than those achieved by other methods. However, one has to take into consideration that a reliable reference protein is needed to assess the distribution of the protein of interest.
Fluorescent staining by immunohistochemistry allows us to routinely identify the localization of proteins across brain areas as well as within different neuronal compartments. To identify the different compartments, specific markers are used. Typically, antibodies against synapsin and synaptophysin11 can be used to label synaptic vesicles, while antibodies against bassoon label the active zone of a presynaptic terminal12. Vesicular transporters, such as the vesicular glutamate transporters (vGluT) or vesicular GABA transporter (vGAT), are used to label excitatory13 and inhibitory14 presynaptic terminals, respectively. On the postsynaptic side, antibodies against the Homer protein can be employed to mark postsynaptic terminals, and antibodies against postsynaptic density protein 95 (PSD95)15,16,17 or gephyrin18,19,20 can label excitatory or inhibitory postsynaptic terminals, respectively. By using antibodies against a protein of interest and markers such as the ones described above, one can determine the localization of such protein. Many studies to date have done this in a qualitative manner21. However, to reliably determine the differential distribution of a specific synaptic protein, one must not only determine its presence or absence but also its relative concentration. The heterogeneity of sizes and density of synapses makes it important to establish a ratio between the synaptic marker and the protein of interest. Otherwise, synapse-rich regions such as the non-pyramidal layers of the hippocampus and the molecular layer of the cerebellum will show a high density of synaptic proteins, only due to the higher density of synapses but not due to a strong presence of that protein in each synapse (e.g., Wallrafen and Dresbach1). On the other hand, proteins in the neuronal soma (e.g., TGN3822) will usually show strong presence in the hippocampal pyramidal cell layer or hippocampal or cerebellar granule cell layer due to the high concentration of neuronal cell bodies in those areas. Therefore, this non-homogeneous distribution of structures, in this case synapses, can lead to a false estimation of the distribution of the protein of interest itself. Furthermore, there is an intrinsic variability in staining intensities across samples in immunohistochemical stainings. The protocol described here takes this into consideration and avoids such biases, as well as other caveats that arise from immunohistochemical methods.
In our recent study, we have used this method to describe the differential expression of Mover (also called TPRGL23 or SVAP3024) across 16 different brain areas1. Mover is a vertebrate-specific synaptic protein that can be found in association to synaptic vesicles and influences neurotransmitter release25,26,27. We have related the Mover expression to the abundance of synaptic vesicles, by staining for synaptophysin as a synaptic vesicle reference marker. We found high levels of Mover particularly in the septal nuclei, the ventral pallidum, and the amygdala. Within the hippocampus, we found a heterogeneous distribution of Mover, with high levels in the layers associated with intra-hippocampal computation, and low levels in input- and output layers.

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			<td align="right">3123000012</td>
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		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Vortex Genius 3&#160;</td>
			<td>IKA</td>
			<td align="right">3340001</td>
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			<td height="40" style="height:40px;">Menzel microscope slides</td>
			<td>Fisher Scientific</td>
			<td colspan="2">&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; 10144633CF</td>
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			<td height="40" style="height:40px;">Stereoscope</td>
			<td>Leica</td>
			<td></td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;">LSM800</td>
			<td>Zeiss</td>
			<td></td>
			<td>Confocal microscope</td>
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		<tr height="40">
			<td height="40" style="height:40px;">freezing microtome</td>
			<td>Leica</td>
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			<td></td>
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			<td height="40" style="height:40px;width:216px;">PBS (10X)</td>
			<td style="width:167px;">Roche</td>
			<td style="width:187px;">&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; 11666789001</td>
			<td></td>
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			<td height="40" style="height:40px;">PFA</td>
			<td>Sigma</td>
			<td colspan="2">&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; P6148-1kg</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">NaCl</td>
			<td>BioFroxx</td>
			<td>1394KG001</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">sucrose</td>
			<td>neoFroxx</td>
			<td>1104KG001</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Tissue Tek</td>
			<td>Sakura&#160;</td>
			<td align="right">4583</td>
			<td>OCT</td>
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		<tr height="40">
			<td height="40" style="height:40px;">Na2HPO4</td>
			<td>BioFroxx</td>
			<td>5155KG001</td>
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		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">NaH2PO4</td>
			<td>Merck</td>
			<td align="right">1,063,460,500</td>
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			<td height="40" style="height:40px;">normal goat serum</td>
			<td>Merck Millipore</td>
			<td>S26-100ML</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;">normal donkey serum</td>
			<td>Merck</td>
			<td>S30-100ML</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;">Triton X-100</td>
			<td>Merck</td>
			<td align="right">1,086,031,000</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;">rabbit anti-Mover</td>
			<td>Synaptic Systems</td>
			<td>RRID: AB_10804285</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;">guinea pig anti-Synaptophysin</td>
			<td>Synaptic Systems</td>
			<td>RRID: AB_1210382</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">donkey anti-rabbit AF647</td>
			<td>Jackson ImmunoResearch</td>
			<td>RRID: AB_2492288</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">goat anti-mouse AF488</td>
			<td>Jackson ImmunoResearch</td>
			<td>RRID: AB_2337438</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">Mowiol4-88</td>
			<td style="width:167px;">Calbiochem</td>
			<td style="width:187px;">&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; 475904</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">ZEN2 blue software</td>
			<td style="width:167px;">Zeiss</td>
			<td style="width:187px;"></td>
			<td>Microscopy software</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">FIJI</td>
			<td style="width:167px;">ImageJ</td>
			<td style="width:187px;"></td>
			<td>Analysis software</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Microsoft Excel</td>
			<td>Microsoft</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

quantifying the heterogeneous distribution of a synaptic protein in the mouse brain using immunofluorescence

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a protein, it is vital to also determine its distribution. Here, we describe a protocol employing immunofluorescence, confocal microscopy, and computer-based analysis to determine the distribution of the synaptic protein Mover (also called TPRGL or SVAP30). We compare the distribution of Mover to that of the synaptic vesicle protein synaptophysin, thereby determining the distribution of Mover in a quantitative manner relative to the abundance of synaptic vesicles. Notably, this method could potentially be implemented to allow for comparison of the distribution of proteins using different antibodies or microscopes or across different studies. Our method circumvents the inherent variability of immunofluorescent stainings by yielding a ratio rather than absolute fluorescence levels. Additionally, the method we describe enables the researcher to analyze the distribution of a protein on different levels: from whole brain slices to brain regions to different subregions in one brain area, such as the different layers of the hippocampus or sensory cortices. Mover is a vertebrate-specific protein that is associated with synaptic vesicles. With this method, we show that Mover is heterogeneously distributed across brain areas, with high levels in the ventral pallidum, the septal nuclei, and the amygdala, and also within single brain areas, such as the different layers of the hippocampus.

Representative staining patterns of different markers can be seen in Figure 1. The pattern varies depending on the distribution of the protein. Examples of five rostro-caudal levels are shown in columns (A)-(E). A representative DAPI staining is shown in the first row: DAPI adheres to the DNA of a cell and thus nuclei are stained. This results in a punctate pattern. Regions with a high cell density are brighter than regions w...

Watch this Scientific Journal Video about Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence at JoVE.com

Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

Here, we describe a quantitative approach to determining the distribution of a synaptic protein relative to a marker protein using immunofluorescence staining, confocal microscopy, and computer-based analysis.

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a ...

quantifying-heterogeneous-distribution-synaptic-protein-mouse-brain

Research

JoVE Journal

Neuroscience

7.8K Views.  University Medical Center Göttingen. Here, we describe a quantitative approach to determining the distribution of a synaptic protein relative to a marker protein using immunofluorescence staining, confocal microscopy, and computer-based analysis.

Video: Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to cis-regulatory elements (CREs), short (&tilde;300-600 bp) stretches of genomic DNA which can lie upstream, downstream, or within the introns of the genes they control. CREs (i.e., enhancers/promoters) typically consist of multiple clustered binding sites for both transcriptional activators and repressors1-3. They serve as logical integrators of transcriptional input giving a unitary output in the form of spatiotemporally precise and quantitatively exact promoter activity. Most studies of mammalian cis-regulation to date have relied on mouse transgenesis as a means of assaying the enhancer function of CREs4-5. This technique is time-consuming, costly and, on account of insertion site effects, largely non-quantitative. On the other hand, quantitative assays for mammalian CRE function have been developed in tissue culture systems (e.g., dual luciferase assays), but the in vivo relevance of these results is often uncertain. 
Electroporation offers an excellent alternative to traditional mouse transgenesis in that it permits both spatiotemporal and quantitative assessment of cis-regulatory activity in living mammalian tissue. This technique has been particularly useful in the analysis of cis-regulation in the central nervous system, especially in the cerebral cortex and the retina6-8. While mouse retinal electroporation, both in vivo and ex vivo, has been developed and extensively described by Matsuda and Cepko6-7,9, we have recently developed a simple approach to quantify the activity of photoreceptor-specific CREs in electroporated mouse retinas10. Given that the amount of DNA that is introduced into the retina by electroporation can vary from experiment to experiment, it is necessary to include a co-electroporated 'loading control' in all experiments. In this respect, the technique is very similar to the dual luciferase assay used to quantify promoter activity in cultured cells. 
When assaying photoreceptor cis-regulatory activity, electroporation is usually performed in newborn mice (postnatal day 0, P0) which is the time of peak rod production11-12. Once retinal cell types become post-mitotic, electroporation is much less efficient. Given the high rate of rod birth in newborn mice and the fact that rods constitute more than 70% of the cells in the adult mouse retina, the majority of cells that are electroporated at P0 are rods. For this reason, rod photoreceptors are the easiest retinal cell type to study via electroporation. The technique we describe here is primarily useful for quantifying the activity of photoreceptor CREs.

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

This protocol describes a simple and inexpensive way to quantify the activity of cis-regulatory elements (i.e., enhancer/promoters) in living mouse retinas via explant electroporation. DNA preparation, retinal dissection, electroporation, retinal explant culture, and post-fixation analysis and quantification are described.

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to ...

Monitoring Changes in the Intracellular Calcium Concentration and Synaptic Efficacy in the Mollusc Aplysia

It has been suggested that changes in intracellular calcium mediate the induction of a number of important forms of synaptic plasticity (e.g., homosynaptic facilitation) 1. These hypotheses can be tested by simultaneously monitoring changes in intracellular calcium and alterations in synaptic efficacy. We demonstrate how this can be accomplished by combining calcium imaging with intracellular recording techniques. Our experiments are conducted in a buccal ganglion of the mollusc Aplysia californica. This preparation has a number of experimentally advantageous features: Ganglia can be easily removed from Aplysia and experiments use adult neurons that make normal synaptic connections and have a normal ion channel distribution. Due to the low metabolic rate of the animal and the relatively low temperatures (14-16 &deg;C) that are natural for Aplysia, preparations are stable for long periods of time. 
To detect changes in intracellular free calcium we will use the cell impermeant version of Calcium Orange 2 which is easily 'loaded' into a neuron via iontophoresis. When this long wavelength fluorescent dye binds to calcium, fluorescence intensity increases. Calcium Orange has fast kinetic properties 3 and, unlike ratiometric dyes (e.g., Fura 2), requires no filter wheel for imaging. It is fairly photo stable and less phototoxic than other dyes (e.g., fluo-3) 2,4. Like all non-ratiometric dyes, Calcium Orange indicates relative changes in calcium concentration. But, because it is not possible to account for changes in dye concentration due to loading and diffusion, it can not be calibrated to provide absolute calcium concentrations. 
An upright, fixed stage, compound microscope was used to image neurons with a CCD camera capable of recording around 30 frames per second. In Aplysia this temporal resolution is more than adequate to detect even a single spike induced alteration in the intracellular calcium concentration. Sharp electrodes are simultaneously used to induce and record synaptic transmission in identified pre- and postsynaptic neurons. At the conclusion of each trial, a custom script combines electrophysiology and imaging data. To ensure proper synchronization we use a light pulse from a LED mounted in the camera port of the microscope. Manipulation of presynaptic calcium levels (e.g. via intracellular EGTA injection) allows us to test specific hypotheses, concerning the role of intracellular calcium in mediating various forms of plasticity.

Monitoring Changes in the Intracellular Calcium Concentration and Synaptic Efficacy in the Mollusc Aplysia

We demonstrate how changes in the intracellular free calcium concentration and synaptic efficacy can be simultaneously monitored in a ganglion preparation of Aplysia. We image intracellular calcium using a fluorescent dye, Calcium Orange, and induce and monitor synaptic transmission with sharp (intracellular) electrodes.

It has been suggested that changes in intracellular calcium mediate the induction of a number of important forms of synaptic plasticity (e.g., ...

Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

Ganglion cells are the output neurons of the retina and their activity reflects the integration of multiple synaptic inputs arising from specific neural circuits. Patch clamp techniques, in voltage clamp and current clamp configurations, are commonly used to study the physiological properties of neurons and to characterize their synaptic inputs. Although the application of these techniques is highly informative, they pose various limitations. For example, it is difficult to quantify how the precise interactions of excitatory and inhibitory inputs determine response output. To address this issue, we used a modified current clamp technique, dynamic clamp, also called conductance clamp 1, 2, 3 and examined the impact of excitatory and inhibitory synaptic inputs on neuronal excitability. This technique requires the injection of current into the cell and is dependent on the real-time feedback of its membrane potential at that time. The injected current is calculated from predetermined excitatory and inhibitory synaptic conductances, their reversal potentials and the cell's instantaneous membrane potential. Details on the experimental procedures, patch clamping cells to achieve a whole-cell configuration and employment of the dynamic clamp technique are illustrated in this video article. Here, we show the responses of mouse retinal ganglion cells to various conductance waveforms obtained from physiological experiments in control conditions or in the presence of drugs. Furthermore, we show the use of artificial excitatory and inhibitory conductances generated using alpha functions to investigate the responses of the cells.

This video article illustrates the set-up, the procedures to patch cell bodies and how to implement dynamic clamp recordings from ganglion cells in whole-mount mouse retinae. This technique allows the investigation of the precise contribution of excitatory and inhibitory synaptic inputs, and their relative magnitude and timing to neuronal spiking.

Ganglion cells are the output neurons of the  retina and their activity reflects the integration of multiple synaptic inputs  arising from specific neural ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

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 ...

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We ...

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 ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

The ubiquitin-proteasome system is a key regulator of protein degradation and a variety of other cellular processes in eukaryotes. In the brain, increases in ubiquitin-proteasome activity are critical for synaptic plasticity and memory formation and aberrant changes in this system are associated with a variety of neurological, neurodegenerative and psychiatric disorders. One of the issues in studying ubiquitin-proteasome functioning in the brain is that it is present in all cellular compartments, in which the protein targets, functional role and mechanisms of regulation can vary widely. As a result, the ability to directly compare brain ubiquitin protein targeting and proteasome catalytic activity in different subcellular compartments within the same animal is critical for fully understanding how the UPS contributes to synaptic plasticity, memory and disease. The method described here allows collection of nuclear, cytoplasmic and crude synaptic fractions from the same rodent (rat) brain, followed by simultaneous quantification of proteasome catalytic activity (indirectly, providing activity of the proteasome core only) and linkage-specific ubiquitin protein tagging. Thus, the method can be used to directly compare subcellular changes in ubiquitin-proteasome activity in different brain regions in the same animal during synaptic plasticity, memory formation and different disease states. This method can also be used to assess the subcellular distribution and function of other proteins within the same animal.

This protocol is designed to efficiently quantify ubiquitin-proteasome system (UPS) activity in different cellular compartments of the rodent brain. Users are able to examine UPS functioning in nuclear, cytoplasmic and synaptic fractions in the same animal, reducing the amount of time and number of animals needed to perform these complex analyses.

The ubiquitin-proteasome system is a key regulator of protein degradation and a variety of other cellular processes in eukaryotes. In the brain, increases ...