Name: quantifying subcellular ubiquitin-proteasome activity in the rodent brain
Uploaded: 2019-05-21T14:30:00
Description: Watch this Scientific Journal Video about Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain at JoVE.com

This work was supported by startup funds from the College of Agricultural and Life Sciences and the College of Science at Virginia Tech. T.M. is supported by the George Washington Carver Program at Virginia Tech.

All procedures including animal subjects have been approved by the Virginia Polytechnic Institute and State University Institutional Animal Care and Use Committee (IACUC).
1. Collection and Dissection of Rodent Brain Tissue
NOTE: This protocol can be applied to a variety of brain regions and used with various tissue collection procedures. Below is the procedure used in our lab for subcellular of rat brain tissue, using 8-9 week old male Sprague Dawley...

<ol>
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Here, we demonstrate an efficient method for quantifying changes in ubiquitin-proteasome activity across different subcellular compartments in the same animal. Currently, most attempts at measuring subcellular changes in activity of the ubiquitin-proteasome system have been limited to a single compartment per sample, resulting in the need to repeat experiments. This leads to significant costs and loss of animal life. Our protocol alleviates this problem by splitting hemispheres, allowing different cellular fractions to b...

The ubiquitin-proteasome system (UPS) is a complex network of interconnected protein structures and ligases that controls the degradation of most short-lived proteins in cells1. In this system, proteins are marked for degradation or other cellular processes/fates by the small modifier ubiquitin. A target protein can acquire 1-7 ubiquitin modifications, which can link together at one of seven lysine (K) sites (K6, K11, K27, K29, K33, K48 and K63) or the N-terminal methionine (M1; as known as linear) in the previous ubiquitin2. Some of these polyubiquitin tags are degradation-specific (K48)3, while others are largely independent of the protein degradation process (M1)4,5,6. Thus, the protein ubiquitination process is incredibly complex and the ability to quantify changes in a specific polyubiquitin tag is critical for ultimately understanding the role of that given modification in cellular functioning. Further complicating the study of this system, the proteasome, which is the catalytic structure of the UPS7, both degrades proteins but can also be involved in other non-proteolytic processes8,9. Not surprisingly then, since its initial discovery, normal and aberrant ubiquitin-proteasome activity has been implicated in long-term memory formation and a variety of disease states, including many neurological, neurodegenerative and psychiatric disorders10,11. As a result, methods which can effectively and efficiently quantify UPS activity in the brain are critical for ultimately understanding how this system is dysregulated in disease states and the eventual development of treatment options targeting ubiquitin and/or proteasome functioning.
There are a number of issues in quantifying ubiquitin-proteasome activity in brain tissue from rats and mice, which are the most common model systems used to study UPS function, including 1) the diversity of ubiquitin modifications, and 2) distribution and differential regulation of UPS functioning across subcellular compartments12,13,14. For example, many of the early demonstrations of ubiquitin-proteasome function in the brain during memory formation used whole cell lysates and indicated time-dependent increases in both protein ubiquitination and proteasome activity15,16,17,18,19,20. However, we recently found that ubiquitin-proteasome activity varied widely across subcellular compartments in response to learning, with simultaneous increases in some regions and decreases in others, a pattern that differs significantly from what was previously reported in whole cell lysates21. This is consistent with the limitation of a whole cell approach, as it cannot dissociate the contribution of changes in UPS activity across different subcellular compartments. Though more recent studies have employed synaptic fraction protocols to study the UPS specifically at synapses in response to learning22,23,24, the methods used occlude the ability to measure nuclear and cytoplasmic ubiquitin-proteasome changes in the same animal. This results in an unnecessary need to repeat experiments multiple times, collecting a different subcellular fraction in each. Not only does this result in a greater loss of animal lives, but it eliminates the ability to directly compare UPS activity across different subcellular compartments in response to a given event or during a specific disease state. Considering that protein targets of ubiquitin and the proteasome vary widely throughout the cell, understanding how ubiquitin-proteasome signaling differs in distinct subcellular compartments is critical for identifying the functional role of the UPS in the brain during memory formation and neurological, neurodegenerative and psychiatric disorders.
To address this need, we recently developed a procedure in which nuclear, cytoplasmic and synaptic fractions could be collected for a given brain region from the same animal21. Additionally, to account for the limited amount of protein that can be obtained from collecting multiple subcellular fractions from the same sample, we optimized previously established protocols to assay in vitro&#160;proteasome activity and linkage-specific protein ubiquitination in lysed cells collected from rodent brain tissue. Using this protocol, we were able to collect and directly compare learning-dependent changes in proteasome activity, K48, K63, M1 and overall protein polyubiquitination levels in the nucleus and cytoplasm and at synapses in the lateral amygdala of rats. Here, we describe in detail our procedure (Figure 1), which could significantly improve our understanding of how the UPS is involved in long-term memory formation and various disease states. However, it should be noted that the in vitro proteasome activity discussed in our protocol, while widely used, does not directly measure the activity of complete 26S proteasome complexes. Rather, this assay measures the activity of the 20S core, meaning it can only serve as a proxy to understand the activity of the core itself as opposed to the entire 26S proteasome complex.

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			<td height="40" style="height:40px;width:222px;">0.5M EDTA</td>
			<td style="width:143px;">Fisher</td>
			<td style="width:259px;">15575020</td>
			<td style="width:255px;">Various other vendors</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">20S Proteasome Activity Kit</td>
			<td>Millipore Sigma</td>
			<td>APT280</td>
			<td>Other vendors carry different versions</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:222px;">ATP</td>
			<td style="width:143px;">Fisher</td>
			<td style="width:259px;">FERR1441</td>
			<td style="width:255px;">Various other vendors</td>
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			<td>Cell signaling</td>
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			<td>Various other vendors</td>
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			<td height="40" style="height:40px;">Beta-tubulin antibody</td>
			<td>Cell signaling</td>
			<td>2128T</td>
			<td>Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;">BioTek Synergy H1 plate reader</td>
			<td>BioTek</td>
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			<td>Other vendors carry different versions</td>
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			<td height="40" style="height:40px;">clasto lactacystin b-lactone</td>
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			<td>L7035</td>
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			<td>Fisher</td>
			<td>033377B</td>
			<td>Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;">DMSO</td>
			<td>DMSO</td>
			<td>D8418</td>
			<td>Varous other vendors</td>
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			<td height="40" style="height:40px;width:222px;">DTT</td>
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			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">G5516</td>
			<td style="width:255px;">Various other vendors</td>
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			<td height="40" style="height:40px;">H3 antibody</td>
			<td>Abcam</td>
			<td>ab1791</td>
			<td>Various other vendors</td>
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			<td height="40" style="height:40px;width:222px;">HEPES</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">H3375</td>
			<td style="width:255px;">Various other vendors</td>
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			<td height="40" style="height:40px;">Hydrochloric acid</td>
			<td>Fisher</td>
			<td>SA48</td>
			<td>Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;width:222px;">IGEPAL (NP-40)</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">I3021</td>
			<td style="width:255px;">Various other vendors</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">K48 Ubiquitin Antibody</td>
			<td>Abcam</td>
			<td>ab140601</td>
			<td>Various other vendors</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">K63 Ubiquitin Antibody</td>
			<td>Abcam</td>
			<td>ab179434</td>
			<td>Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;width:222px;">KCl</td>
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			<td>Life Sensors</td>
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			<td>Only M1 antibody</td>
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			<td>Carried by Millipore-Sigma</td>
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			<td height="40" style="height:40px;width:222px;">NaCl</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">S3014</td>
			<td style="width:255px;">Various other vendors</td>
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			<td height="40" style="height:40px;width:222px;">Phosphatase Inhibitor</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">524625</td>
			<td style="width:255px;">Various other vendors</td>
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			<td>Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;width:222px;">Protease Inhibitor</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">P8340</td>
			<td style="width:255px;">Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;">PSD95 antibody</td>
			<td>Cell signaling</td>
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			<td>Various other vendors</td>
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			<td height="50" style="height:50px;width:222px;">SDS</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">L3771</td>
			<td style="width:255px;">Various other vendors</td>
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			<td height="45" style="height:46px;">Sodium hydroxide</td>
			<td>Fisher</td>
			<td>SS255</td>
			<td>Various other vendors</td>
		</tr>
		<tr height="48">
			<td height="48" style="height:48px;width:222px;">Sucrose</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">S0389</td>
			<td style="width:255px;">Various other vendors</td>
		</tr>
		<tr height="58">
			<td height="58" style="height:58px;">TBS</td>
			<td>Alfa Aesar</td>
			<td>J62938</td>
			<td>Varous other vendors</td>
		</tr>
		<tr height="49">
			<td height="49" style="height:50px;width:222px;">Tris</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">T1503</td>
			<td style="width:255px;">Various other vendors</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Tween-20</td>
			<td>Fisher</td>
			<td>BP337-100</td>
			<td>Various other vendors</td>
		</tr>
		<tr height="48">
			<td height="48" style="height:48px;">Ubiquitin Antibody</td>
			<td>Enzo Life Sciences</td>
			<td>BML-PW8810</td>
			<td>Various other vendors</td>
		</tr>
	</tbody>
</table>

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.

Using the procedure described here, nuclear, cytoplasmic and synaptic fractions were collected from the lateral amygdala of the rat brain (Figure 1). Purity of the individual fractions were confirmed via Western blotting, probing with antibodies against proteins that should be enriched or depleted in the lysate. In the first hemisphere where a crude synaptic fraction was collected, postsynaptic density protein 95 (PSD95) was present in the synaptic, but not n...

Watch this Scientific Journal Video about Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain at JoVE.com

Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

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 ...

This protocol measures subcellular protein ubiquitination levels and proteasome activity in the rodent brain allowing within subjects comparisons of how ubiquitin-proteasome activity changes in response to cellular activity, learning, or disease. The protocol allows collection of synaptic, cytoplasmic, and nuclear fractions from the same rodent brain. Minimizing loss, it could be performed with small tissue quantities and basic laboratory equipment.Demonstrating the procedure will be Taylor McFadden, a graduate student from my laboratory. Begin this procedure with collection and dissection of rodent brain tissue as described in the text protocol. Ensure that the hemisphere used is counter balanced across extraction conditions for each experimental group.Remove the 1.5 milliliter centrifuge microtube containing one hemisphere of the region of interest from the minus 80 degree Celsius freezer. Using a scalpel, transfer the frozen brain tissue to a two milliliter glass Teflon homogenizer. Add 500 microliters of lysis buffer to the Teflon tube.Using pestle B, homogenize the same tissue with 15 strokes until no visible amount of solid material is present. Use a turning motion during each stroke. With a 1, 000 microliter pipette, transfer the homogenized sample to a new 1.5 milliliter microcentrifuge tube.Place the tube on wet ice and incubate for 30 minutes. Place the tube in the microcentrifuge and spin for 10 minutes at 845 times g in four degrees Celsius. After completion, carefully remove the supernatant by pipetting and place into a new 1.5 milliliter microcentrifuge tube.This is the cytoplasmic fraction. Add 50 microliters of extraction buffer to the resulting pellet and resuspend by pipetting. Do not vortex the pellet.Place the tube containing the resuspended pellet on ice and incubate for 30 minutes. Then centrifuge the tube for 20 minutes at 21, 456 times g in four degrees Celsius. Following centrifugation, carefully remove the supernatant by pipetting and place into a new 1.5 milliliter microcentrifuge tube.This is the nuclear fraction. Homogenize one hemisphere of the region of interest as before except to use 500 microliters of TEVP buffer instead of lysis buffer. Using a 1, 000 microliter pipette, transfer the homogenized sample to a new 1.5 milliliter microcentrifuge tube.Centrifuge the sample at 1, 000 times g for 10 minutes at four degrees Celsius. Collect the supernatant and transfer it to a new 1.5 milliliter microcentrifuge tube using a 1, 000 microliter pipette. Centrifuge the sample at 10, 000 times g for 10 minutes at four degrees Celsius.Discard the original pellet which contains nuclei and the large debris. Transfer the supernatant to a new 1.5 milliliter microcentrifuge tube. This is the cytosolic fraction.Add 50 microliters of homogenization buffer to the pellet and resuspend by pipetting until no solid material is visible. Centrifuge the sample at 20, 000 times g for 10 minutes at four degrees Celsius. Transfer the supernatant to a new 1.5 milliliter microcentrifuge tube.This is the crude synaptosomal membrane fraction. To set up the assay, prewarm the plate reader to 37 degrees Celsius and hold through the run. Set the excitation to 360 nanometers and emission to 460 nanometers.If the 96 well plate used is clear, set the optics position to bottom. If a dark 96 well plate is used, set optics position to top. Program a kinetic run with a time of two hours scanning every 30 minutes.Reconstitute the 20S 10X assay buffer provided in the kit with 13.5 milliliters of ultrapure water. Add 14 microliters of 100 millimolar ATP to the now 1X buffer. This significantly enhances proteasome activity in the samples and improves assay reliability.Reconstitute the AMC standard provided in the kit with 100 microliters of DMSO. Perform steps using the AMC standard in the dark or under low light conditions as the standard is light sensitive. Create a standard curve of AMC from a series of high to low AMC concentrations.This curve will be used for plate reader calibration and analysis of proteasome activity in the homogenized samples. Reconstitute the proteasome subtrate provided in the kit with 65 microliters of DMSO. Also perform this step in the dark or under low light conditions as the substrate is light sensitive.Then create a one to 20 dilution of the proteasome substrate in a new 1.5 milliliter microcentrifuge tube using 20S assay buffer. Add a normalized amount of the desired samples to a 96 well plate. Run each sample in duplicate.The amount of sample needed varies based on tissue preparation. Generally, 10 to 20 micrograms is sufficient for any subcellular fraction. Bring the sample well volume to 80 microliters with ultrapure water.The amount added depends on the volume of sample added. In two separate wells, add 80 microliters of water alone. These will be the assay blanks.Add 10 microliters of 20S assay buffer to each well including assay blanks. Use a repeater or automated pipette to ensure consistent assay volume across the wells. Turn off the lights or enter a dark room.Add all 100 microliters of diluted AMC standards to a new well using a single well for each standard. In the dark, use a repeater or automated pipette to add 10 microliters of diluted proteasome substrate to wells containing sample and assay blanks but not the AMC standard. Place the plate into the plate reader and start the kinetic run.Perform quantification of diverse polyubiquitin tags in different subcellular fractions collected from rodent brain tissue using a variety of standard Western blotting protocols in combination with unique linkage-specific polyubiquitin antibodies. Some ubiquitin Western blot images will provide columns of distinct bands while others produces smear-like pattern with few or no clear lines. For quantification of imaged ubiquitin Western blots, draw a box around the column that extends the entire molecular standards ladder.Adjust the box if ubiquitin staining does extend through the entire ladder. This is common for lysine 48 modifications and varies widely across subcellular compartments. Finally, subtract out the background which is calculated as the mean optical density of the background immediately surrounding the column on all sides.Shown here is the quantification of proteasome activity in different fractions collected from the lateral amygdala of the same animal. During the in vitro proteasome activity assay, relative fluorescent units detected increased from the beginning to the end of the assay in the synaptic fraction, the cytoplasmic fraction, and the nuclear fraction. The proteasome inhibitor beta-lac prevented RFUs from changing across the session.Subcellular differences were observed in proteasome activity in the lateral amygdala of the same animal. An increase in nuclear proteasome activity was detected in trained animals relative to controls which corresponded to a decrease in activity within the synaptic fraction. Cytoplasmic proteasome activity remained at baseline.Shown here are subcellular differences in linkage-specific protein ubiquitination in the lateral amygdala of the same animal following learning. There was an increase in overall ubiquitination in the nuclear fraction following learning which correlated with a decrease in the cytoplasmic fraction. Following learning, linear ubiquitination increased in the nuclear fraction but not cytoplasmic or synaptic fractions.Interestingly, K63 ubiquitination increased in the nuclear fraction following learning which correlated with a decrease in the synaptic fraction. Whereas K48 uniquitination increased in the nuclear and cytoplasmic fractions following learning but not in the synaptic fraction. This protocol can also be used to understand the subcellular distribution and function of other proteins within the same animal.

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Simultaneous fMRI and Electrophysiology in the Rodent Brain

To examine the neural basis of the blood oxygenation level dependent (BOLD) magnetic resonance imaging (MRI) signal, we have developed a rodent model in which functional MRI data and in vivo intracortical recording can be performed simultaneously. The combination of MRI and electrical recording is technically challenging because the electrodes used for recording distort the MRI images and the MRI acquisition induces noise in the electrical recording. To minimize the mutual interference of the two modalities, glass microelectrodes were used rather than metal and a noise removal algorithm was implemented for the electrophysiology data. In our studies, two microelectrodes were separately implanted in bilateral primary somatosensory cortices (SI) of the rat and fixed in place. One coronal slice covering the electrode tips was selected for functional MRI. Electrode shafts and fixation positions were not included in the image slice to avoid imaging artifacts. The removed scalp was replaced with toothpaste to reduce susceptibility mismatch and prevent Gibbs ringing artifacts in the images. The artifact structure induced in the electrical recordings by the rapidly-switching magnetic fields during image acquisition was characterized by averaging all cycles of scans for each run. The noise structure during imaging was then subtracted from original recordings. The denoised time courses were then used for further analysis in combination with the fMRI data. As an example, the simultaneous acquisition was used to determine the relationship between spontaneous fMRI BOLD signals and band-limited intracortical electrical activity. Simultaneous fMRI and electrophysiological recording in the rodent will provide a platform for many exciting applications in neuroscience in addition to elucidating the relationship between the fMRI BOLD signal and neuronal activity.

We have developed a method for simultaneous functional magnetic resonance imaging and electrophysiological recording in the rodent brain, providing a platform for the investigation of the relationship between neural activity and the blood oxygenation level dependent (BOLD) MRI signal.

To examine the neural basis of the blood oxygenation level dependent (BOLD) magnetic resonance imaging (MRI) signal, we have developed a rodent model in ...

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 ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

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

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

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

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

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

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Personalized Needles for Microinjections in the Rodent Brain

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to deliver gene or cell therapy products. Unfortunately, current microinjection techniques use steel or glass needles that are suboptimal for multiple reasons: in particular, steel needles may cause tissue damage, and glass needles may bend when lowered deeply into the brain, missing the target region. In this article, we describe a protocol to prepare and use quartz needles that combine a number of useful features. These needles do not produce detectable tissue damage and, being very rigid, ensure reliable delivery in the desired brain region even when using deep coordinates. Moreover, it is possible to personalize the design of the needle by making multiple holes of the desired diameter. Multiple holes facilitate the injection of large amounts of solution within a larger area, whereas large holes facilitate the injection of cells. In addition, these quartz needles can be cleaned and re-used, such that the procedure becomes cost-effective.

We describe here a protocol for microinjection in the rodent brain that uses quartz needles. These needles do not produce detectable tissue damage and ensure reliable delivery even in deep regions. Moreover, they can be adapted to research needs by personalized designs and can be re-used.

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is tightly regulated during development and neuronal maturation. Importantly, deficits in synapse number can lead to cognitive dysfunction. Therefore, the evaluation of synapse number is an integral part of neurobiology. However, as synapses are small and highly compact in the intact brain, the assessment of absolute number is challenging. This protocol describes a method to easily identify and evaluate synapses in hippocampal rodent slices using immunofluorescence microscopy. It includes a three-step procedure to evaluate synapses in high-quality confocal microscopy images by analyzing the co-localization of pre- and postsynaptic proteins in hippocampal slices. It also explains how the analysis is performed and gives representative examples from both excitatory and inhibitory synapses. This protocol provides a solid foundation for the analysis of synapses and can be applied to any research investigating the structure and function of the brain.

A protocol for accurately identifying and analyzing synapses in hippocampal slices using immunofluorescence is outlined in this article.

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is ...

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) is a promising approach when pharmacological treatment fails or neurosurgery is not recommended. Acute brain slices coupled to microelectrode arrays (MEAs) represent a valuable tool to study neuronal network interactions and their modulation by electrical stimulation. As compared to conventional extracellular recording techniques, they provide the added advantages of a greater number of observation points and a known inter-electrode distance, which allow studying the propagation path and speed of electrophysiological signals. However, tissue oxygenation may be greatly impaired during MEA recording, requiring a high perfusion rate, which comes at the cost of decreased signal-to-noise ratio and higher oscillations in the experimental temperature. Electrical stimulation further stresses the brain tissue, making it difficult to pursue prolonged recording/stimulation epochs. Moreover, electrical modulation of brain slice activity needs to target specific structures/pathways within the brain slice, requiring that electrode mapping be easily and quickly performed live during the experiment. Here, we illustrate how to perform the recording and electrical modulation of 4-aminopyridine (4AP)-induced epileptiform activity in rodent brain slices using planar MEAs. We show that the brain tissue obtained from mice outperforms rat brain tissue and is thus better suited for MEA experiments. This protocol guarantees the generation and maintenance of a stable epileptiform pattern that faithfully reproduces the electrophysiological features observed with conventional field potential recording, persists for several hours, and outlasts sustained electrical stimulation for prolonged epochs. Tissue viability throughout the experiment is achieved thanks to the use of a small-volume custom recording chamber allowing for laminar flow and quick solution exchange even at low (1 mL/min) perfusion rates. Quick MEA mapping for real-time monitoring and selection of stimulating electrodes is performed by a custom graphic user interface (GUI).

We illustrate how to perform recording and electrical modulation of 4-aminopyridine-induced epileptiform activity in rodent brain slices using microelectrode arrays. A custom recording chamber maintains tissue viability throughout prolonged experimental sessions. Live electrode mapping and selection of stimulating pairs are performed by a custom graphical user interface.

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) ...

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.

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 ...