This work was funded by NIH grants DA15169 and DA035224 to H.E.M.

All animal handling and tissue harvesting was performed in accordance with the guidelines of the University of Massachusetts Medical School Institutional Animal Care Use Committee (IACUC), following the approved protocol #A1506 (Melikian, P.I.).
Required solutions
Artificial cerebrospinal fluid (ACSF) - Make fresh daily
125 mM NaCl, 2.5 mM KCl, 1.2 mM NaH2PO4, 1.2 mM MgCl2, 2.4 mM CaCl2, 26 mM NaHCO3, and 11 mM glucose
Note: Prepare ACSF as a 10x stock solution, excluding NaHCO3 and glucose. Make 1x working solutions daily from the 10x stock, supplementing with fresh NaHCO3 and glucose.
Sucrose-supplemented ACSF (SACSF) &#8211; Make fresh daily
250 mM sucrose, 2.5 mM KCl, 1.2 mM NaH2PO4, 1.2 mM MgCl2, 2.4 mM CaCl2, 26 mM NaHCO3, and 11 mM glucose
Note: Prepare SACSF as a 10x stock solution, excluding NaHCO3 and glucose. Make 1x working solutions daily from the 10x stock, supplementing with fresh NaHCO3 and glucose.
Sulfo-N-hydroxysuccinyl-SS-Biotin (sulfo-NHS-SS-biotin, Pierce Chemical Company)
Stock solutions should be 200 mg/ml in DMSO and are resistant to multiple freeze/thaw cycles. Aliquots are stored at -20 &#176;C. The succinyl ester is rapidly hydrolyzed in aqueous solution, so working solutions should be prepared immediately prior to applying to slices.
Slice Quench Solution
ACSF supplemented with 100 mM glycine
RIPA Lysis Buffer
10 mM Tris, pH 7.4, 150 mM NaCl, 1.0 mM EDTA, 1% Triton-X-100, 0.1% SDS, 1% Na deoxycholate
RIPA with Protease Inhibitors (RIPA/PI) &#8211; Make fresh daily
RIPA supplemented with 1 &#956;M leupeptin, 1 &#956;M pepstatin, 1 &#956;M aprotinin, and 1 mM phenylmethyl sulfonyl fluoride.
1. Prepare Brain Slices
<ol>
	<li>Make fresh 1x SACSF and 1x ACSF</li>
	<li>Chill SACSF on ice in a small beaker. This will be used to hold the freshly harvested mouse brain.</li>
	<li>Saturate the ACSF and SACSF with oxygen by bubbling with 95%/5% O2/CO2, 20 min on ice.</li>
	<li>P30-38 mice should be used for optimal tissue viability. Sacrifice animals by cervical dislocation and decapitation, and rapidly remove brains into prechilled, oxygenated SACSF.</li>
	<li>Using a vibrating microtome, make 300 &#956;m brain sections in region of interest.</li>
	<li>If desired, slices can be further dissected prior to recovery to enrich for particular brain regions or separate right and left hemispheres to use as control and experimental slices, respectively.</li>
	<li>Using a fire-polished Pasteur pipette, transfer slices to mesh-bottomed chambers set into 24-well plates.</li>
	<li>Allow slices to recover for 40 min, 31 &#176;C in oxygenated ACSF, with continual, gentle bubbling.</li>
</ol>
2. Drug Treatment (if appropriate) and Slice Biotinylation
<ol>
	<li>Following recovery, wash slices 3x in prewarmed (37 &#176;C), oxygenated ACSF bubbling constantly with 95%/5% O2/CO2.</li>
	<li>Add test compounds and incubate with continuous oxygenation.
	<ol>
		<li>For convenience, add a 1/10th volume of 10x concentrated drug, and mix by gently inverting.</li>
		<li>Shake plates gently in a water bath at the desired temperature.</li>
	</ol>
	</li>
	<li>Following drug treatment, rapidly chill slices by washing 3x in ice cold ACSF.</li>
</ol>
3. Biotinylate Surface Proteins
<ol>
	<li>Prepare 1.0 mg/ml sulfo-NHS-SS-biotin in ice cold ACSF immediately prior to labeling.</li>
	<li>Add 0.75 ml sulfo-NHS-SS-biotin to slices and incubate slices on ice, 45 min.</li>
	<li>Wash slices three times quickly with ice cold ACSF, then incubate for 10 min in ice cold ACSF on ice.</li>
	<li>Wash slices three times with ice cold slice quench buffer and incubate with 0.75 ml slice quench buffer two times, 25 min, on ice to quench free sulfo-NHS-SS-biotin.</li>
</ol>
4. Prepare Tissue Lysates
<ol>
	<li>Wash slices three times in ice cold ACSF and transfer each slice to a microcentrifuge tube using a fire-polished Pasteur pipette.</li>
	<li>Gently pellet slice by centrifuging 200 x g, 1 min and carefully aspirate remaining ACSF.</li>
	<li>Add 400 &#181;l ice cold RIPA/PI and break up tissue by pipetting up and down once through a P200 pipette.</li>
	<li>Transfer dissociated slice/RIPA to a fresh tube and incubate 30 min, 4 &#176;C, rotating, to complete lysis.</li>
	<li>Pellet cellular debris by centrifuging, 18,000 x g, 15 min, 4 &#176;C.</li>
	<li>Determine lysate protein concentrations using the BCA protein assay, with bovine serum albumin (BSA) as a standard.</li>
</ol>
5. Isolate Biotinylated Proteins
<ol>
	<li>Optimize Bead/total protein ratio. 
	Note: Individual protein expression levels can vary widely across different brain regions. Unless all of the biotinylated protein in a given amount of tissue lysate is captured, it is not possible to accurately detect any potential changes in surface expression. Therefore, it is imperative to empirically determine the optimal bead/total protein ratio for a particular protein/brain region prior to embarking upon a new slice biotinylation study.
	<ol>
		<li>Incubate 25 &#181;l streptavidin agarose beads with increasing amounts of tissue lysate (approximate range 25-200 &#181;g), and then proceed as described below for binding, washing and elution steps.</li>
		<li>Quantify the resulting immunoreactive bands and choose a bead/lysate ratio in the linear range of binding that will permit accurate quantification of either increased or decreased protein surface expression.</li>
	</ol>
	</li>
	<li>Prepare Streptavidin-Agarose Beads
	<ol>
		<li>Determine total bead volume needed for all samples. Prepare a sufficient bead volume for that amount plus one extra (i.e. 4 samples at&#160;25 &#181;l/sample = 100 &#181;l beads + one extra = 125 &#181;l beads.</li>
		<li>Vortex Streptavidin bead stock and pipette out desired volume. If using a P200 pipette, cut off the end of the tip to prevent blockage or bead damage.</li>
		<li>Wash beads 3 times in 0.5-1.0 ml RIPA/PI to remove preservatives, vortexing after each RIPA addition and collecting beads between washes by centrifuging 18,000 x g, 1 min, at room temperature.</li>
		<li>Aspirate off buffer between washes using a glass Pasteur pipette attached to a vacuum flask. A plastic P200 tip attached to the end of the glass Pasteur pipette provides finer control when aspirating. It is not necessary to remove absolutely all the buffer for the first two&#160;washes, as it risks aspirating beads into the pipette. After the final wash, remove as much excess buffer as possible without aspirating the beads.</li>
		<li>Add RIPA/PI to bring beads back to their original volume, pipetting up and down to resuspend. Avoid vortexing at this point, as beads will stick to the tube wall.</li>
		<li>Aliquot beads into microcentrifuge tubes using a P200 with the tip cut off. Pipette up and down several times between sampling to assure beads remain evenly suspended and dispersed among the tubes.</li>
	</ol>
	</li>
	<li>Bind biotinylated proteins to streptavidin beads
	<ol>
		<li>Distribute cell lysates to the tubes containing the agarose beads. For experiments where multiple samples are being compared, make sure to use the same amount of protein for each sample for accurate comparisons.</li>
		<li>Add additional RIPA/PI to bring to samples to a 200 &#181;l minimal volume. This assures that samples will mix adequately during the incubation and also unifies the protein concentrations across samples.</li>
		<li>Place tubes in a tube rotator and mix overnight at 4 &#176;C.</li>
		<li>In separate tubes, dispense an equivalent of the total lysate volume used for each sample. Alternatively, if the sample volumes are high, a fraction of total lysate volume can be reserved instead, to accommodate maximal load volumes on SDS-PAGE gels. These will be used to normalize the surface expression to the total protein amount.</li>
		<li>Add either an equal volume of 2x or 1/5 volume of 6x SDS-PAGE sample buffer and either incubate at 4 &#176;C, in parallel with bead samples, or store at -20 &#176;C until samples are analyzed.</li>
	</ol>
	</li>
</ol>
6. Elute and Analyze Samples
<ol>
	<li>Pellet beads by centrifuging 18,000 x g, 2 min, at room temperature.</li>
	<li>Aspirate supernatant and wash beads three times with 0.75 ml RIPA. To minimize bead loss, leave a small head volume of buffer above beads between washes. After the final wash, remove as much RIPA as possible, without disrupting the bead pellet.</li>
	<li>Elute biotinylated proteins from streptavidin beads by reducing the disulfide linkage. Add 25 &#181;l 2x SDS-PAGE reducing sample buffer, vortex well and rotate samples 30 min, room temperature. 
	Note: Many membrane proteins have a high tendency to aggregate when boiled in SDS-PAGE sample buffer, severely impairing their electrophoretic mobility. If this is the case for the protein being investigated, avoid heating samples and, instead, elute slowly at room temperature. If boiling is absolutely necessary (e.g. if another protein assessed in parallel requires boiling), sample buffer can be supplemented with 2 M urea (final concentration) to minimize aggregation. While effective in reducing aggregation in some cases, urea often compromises band appearances.</li>
	<li>Analyze samples by immunoblot.</li>
	<li>Thaw total lysate samples and rotate in parallel with bead samples, 30 min, room temperature.</li>
	<li>Separate proteins on SDS-PAGE gels.</li>
	<li>Identify protein(s) of interest by immunoblotting.
	<ol>
		<li>Be certain that bands are detected in the linear range of detection for proper quantification.</li>
		<li>To assure that the biotinylation reagent has not gained access to intracellular proteins via damaged/compromised cells. This is best accomplished by immunoblotting in parallel for an intracellular protein specific to the cell type being investigated.</li>
		<li>Quantify band densities and calculate relative protein surface density as a percent&#160;of the total protein expression level.</li>
	</ol>
	</li>
</ol>

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The authors have no financial interests that conflict with this work.

Despite longstanding knowledge that endocytic trafficking critically impacts synaptic signaling in the brain, it has proved challenging to quantitatively measure changes in protein surface expression in adult neurons.&#160;In this work, we report a reliable approach to label surface protein ex vivo in acute brain slices.&#160;Brain slice preparations have a longstanding history of utility for electrophysiological recordings, as they maintain synaptic connections and cell viability up to hours after their prepara...

Endocytic trafficking is a ubiquitous cellular mechanism that fine-tunes the plasma membrane presentation of a variety of integral membrane proteins.&#160;Endocytosis delivers vital nutrients to the intracellular milieu1 and desensitizes receptor signaling in response to receptor activation2.&#160;Endocytic recycling back to the plasma membrane can additionally enhance cellular signaling by increasing protein expression levels at the cell surface3.&#160;Moreover, membrane trafficking perturbations are implicated in numerous disease and pathological conditions4,5, stressing the need to investigate the molecular mechanisms that govern protein endocytic trafficking.&#160;While many proteins utilize classic clathrin-dependent internalization mechanisms, mounting evidence over the past several years demonstrates that multiple clathrin-independent endocytic mechanisms govern the endocytic potential of an increasing array of proteins6,7.&#160;Thus, the need to investigate endocytic mechanisms facilitating trafficking in physiological relevant systems has grown considerably.
In the brain, endocytic trafficking of receptors, ion channels and neurotransmitter transporters has a primary role in establishing synaptic plasticity8-11 and response to drugs of abuse12-15, ultimately impacting neuronal excitability and synaptic responses.&#160;To date the majority of neuronal trafficking studies rely on either heterologous expression systems or cultured primary neurons, neither of which may reliably reflect mechanisms at play in adult neurons.&#160;Here, we report an approach that uses surface biotinylation to quantitatively measure surface protein levels in acute brain slices derived from adult rodents. Using this approach, we present data that demonstrate that the mouse striatal dopamine transporter rapidly internalizes in response to phorbol ester-mediated protein kinase C (PKC) activation.

<table border="0" cellpadding="0" cellspacing="0" width="682">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:315px;">sulfo NHS-SS-biotin</td>
			<td style="width:83px;">Pierce</td>
			<td style="width:119px;">21331</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:315px;">Streptavidin agarose</td>
			<td style="width:83px;">Pierce</td>
			<td>20347</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:315px;">IgG-free, Protease-free Bovine serum albumin</td>
			<td style="width:83px;">Sigma</td>
			<td>A3059</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:315px;">Vibrating microtome sectioner</td>
			<td style="width:83px;">Various</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:315px;">Shaking water bath</td>
			<td style="width:83px;">various</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:315px;">Milli-cell mesh-bottomed inserts (8&#181;m pore size)</td>
			<td style="width:83px;">Millipore</td>
			<td style="width:119px;">PI8P 012 50</td>
			<td>These can be washed by hand and re-used</td>
		</tr>
	</tbody>
</table>

brain slice biotinylation: an ex vivo approach to measure region-specific plasma membrane protein trafficking in adult neurons

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion channel, and transporter cell surface presentation on a minutes time scale.&#160;There is a broad diversity of mechanisms that control endocytic trafficking of individual proteins. Studies investigating the molecular underpinnings of trafficking have primarily relied upon surface biotinylation to quantitatively measure changes in membrane protein surface expression in response to exogenous stimuli and gene manipulation.&#160;However, this approach has been mainly limited to cultured cells, which may not faithfully reflect the physiologically relevant mechanisms at play in adult neurons.&#160;Moreover, cultured cell approaches may underestimate region-specific differences in trafficking mechanisms.&#160;Here, we describe an approach that extends cell surface biotinylation to the acute brain slice preparation.&#160;We demonstrate that this method provides a high-fidelity approach to measure rapid changes in membrane protein surface levels in adult neurons.&#160;This approach is likely to have broad utility in the field of neuronal endocytic trafficking.

The neuronal dopamine transporter is internalized in response to PKC activation in cell lines16-20. Despite many reports demonstrating PKC-induced DAT surface losses in a variety of cell lines and expression systems, it has been challenging to confirm this finding in cultured dopaminergic neurons21-23. We used mouse striatal slices to directly test whether DAT internalizes in response to PKC activation in adult dopaminergic neurons. Following slice preparation, slices were hemisected along the midli...

Watch this Scientific Journal Video about Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons at JoVE.com

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Neuronal membrane trafficking dynamically controls plasma membrane protein availability and significantly impacts neurotransmission.&#160;To date, it has been challenging to measure neuronal endocytic trafficking in adult neurons.&#160;Here, we describe a highly effective, quantitative method to measure rapid changes in surface protein expression ex vivo in acute brain slices.

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion ...

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12.8K Views.  University of Massachusetts Medical School. Neuronal membrane trafficking dynamically controls plasma membrane protein availability and significantly impacts neurotransmission. To date, it has been challenging to measure neuronal endocytic trafficking in adult neurons. Here, we describe a highly effective, quantitative method to measure rapid changes in surface protein expression ex vivo in acute brain slices.

Video: Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Using an &#945;-Bungarotoxin Binding Site Tag to Study GABA A Receptor Membrane Localization and Trafficking

It is increasingly evident that neurotransmitter receptors, including ionotropic GABA A receptors (GABAAR), exhibit highly dynamic trafficking and cell surface mobility1-7. To study receptor cell surface localization and endocytosis, the technique described here combines the use of fluorescent &#945;-bungarotoxin with cells expressing constructs containing an &#945;-bungarotoxin (Bgt) binding site (BBS). The BBS (WRYYESSLEPYPD) is based on the &#945; subunit of the muscle nicotinic acetylcholine receptor, which binds Bgt with high affinity8,9. Incorporation of the BBS site allows surface localization and measurements of receptor insertion or removal with application of exogenous fluorescent Bgt, as previously described in the tracking of GABAA and metabotropic GABAB receptors2,10. In addition to the BBS site, we inserted a pH-sensitive GFP (pHGFP11) between amino acids 4 and 5 of the mature GABAAR subunit by standard molecular biology and PCR cloning strategies (see Figure 1)12. The BBS is 3&#39; of the pH-sensitive GFP reporter, separated by a 13-amino acid alanine/proline linker. For trafficking studies described in this publication that are based on fixed samples, the pHGFP serves as a reporter of total tagged GABAAR subunit protein levels, allowing normalization of the Bgt labeled receptor population to total receptor population. This minimizes cell to cell Bgt staining signal variability resulting from higher or lower baseline expression of the tagged GABAAR subunits. Furthermore the pHGFP tag enables easy identification of construct expressing cells for live or fixed imaging experiments.

Using an &amp;#945;-Bungarotoxin Binding Site Tag to Study GABA A Receptor Membrane Localization and Trafficking

Here we demonstrate the use of fluorescent Alexa dye coupled to &#945;-bungarotoxin to measure GABA A receptor surface localization and endocytosis in hippocampal neurons. Through the use of constructs bearing a short extracellular tag that binds &#945;-bungarotoxin, analysis of plasma membrane protein endocytic trafficking can be achieved.

It is increasingly evident that neurotransmitter receptors, including ionotropic GABA A receptors (GABAAR), exhibit highly dynamic trafficking and cell ...

Slice It Hot: Acute Adult Brain Slicing in Physiological Temperature

Here we present a protocol for preparation of acute brain slices. This procedure is a critical element for electrophysiological patch-clamp experiments that largely determines the quality of results. It has been shown that omitting the cooling step during cutting procedure is beneficial in obtaining healthy slices and cells, especially when dealing with highly myelinated brain structures from mature animals. Even though the precise mechanism whereby elevated temperature supports neural health can only be speculated upon, it stands to reason that, whenever possible, the temperature in which the slicing is performed should be close to physiological conditions to prevent temperature related artifacts. Another important advantage of this method is the simplicity of the procedure and therefore the short preparation time. In the demonstrated method adult mice are used but the same procedure can be applied with younger mice as well as rats. Also, the following patch clamp experiment is performed on horizontal cerebellar slices, but the same procedure can also be used in other planes as well as other posterior areas of the brain.

In this paper we show a method for preparing acute brain slices in physiological temperature, using a conventional physiological solution without special modifications for the cutting (such as adding sucrose) and without intracardial perfusion of the animal before slice preparation.

Here we present a protocol for preparation of acute brain slices. This procedure is a critical element for electrophysiological patch-clamp experiments ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral tissues. The hypothalamus in the central nervous system (CNS) is the control center for energy and insulin signal response regulation. Chronic inflammation in peripheral tissues and imbalances of certain chemokines (such as CCL5, TNF&#945;, and IL-6) contribute to diabetes and obesity. However, the functional mechanism(s) connecting chemokines and hypothalamic insulin signal regulation still remain unclear.
In vitro primary neuron culture models are convenient and simple models which can be used to investigate insulin signal regulation in hypothalamic neurons. In this study, we introduced exogeneous GLUT4 protein conjugated with GFP (GFP-GLUT4) into primary hypothalamic neurons to track GLUT4 membrane translocation upon insulin stimulation. Time-lapse images of GFP-GLUT4 protein trafficking were recorded by deconvolution microscopy, which allowed users to generate high-speed, high-resolution images without damaging the neurons significantly while conducting the experiment. The contribution of CCR5 in insulin regulated GLUT4 translocation was observed in CCR5 deficient hypothalamic neurons, which were isolated and cultured from CCR5 knockout mice. Our results demonstrated that the GLUT4 membrane translocation efficiency was reduced in CCR5 deficient hypothalamic neurons after insulin stimulation.

This protocol describes a technique for observation of real-time Green Fluorescence Protein (GFP) tagged Glucose Transporter 4 (GLUT4) protein trafficking upon insulin stimulation and characterization of the biological role of CCR5 in the insulin&#8211;GLUT4 signaling pathway with Deconvolution Microscopy.

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral ...

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, very few have been widely accepted. Here, we propose&#160;a new procedure&#160;adapted from the well-known ex vivo brain slice technique, which offers a closer in vivo-like scenario than in vitro preparations, for investigating the early events triggering cell degeneration, as observed in Alzheimer&#39;s disease (AD). This variation consists of simple and easily reproducible steps, which enable preservation of the anatomical cytoarchitecture of the selected brain region and its local functionality in a physiological milieu. Different anatomical areas can be obtained from the same brain, providing the opportunity to perform multiple experiments with the treatments in question in a site-, dose-, and time-dependent manner. Potential limitations&#160;which could affect the outcomes related to this methodology are related to the conservation of the tissue, i.e., the maintenance of its anatomical integrity during the slicing and incubation steps and the section thickness, which can influence the biochemical and immunohistochemical analysis. This approach can be employed for different purposes, such as exploring molecular mechanisms involved in physiological or pathological conditions, drug screening, or dose-response assays. Finally, this protocol could also reduce the number of animals employed in behavioral studies. The application reported here has been recently described and tested for the first time on ex vivo rat brain slices containing the basal forebrain (BF), which is one of the cerebral regions primarily affected in AD. Specifically, it has been demonstrated that the administration of a toxic peptide derived from the C-terminus of acetylcholinesterase (AChE) could prompt an AD-like profile, triggering, along the antero-posterior axis of the BF, a differential expression of proteins altered in AD, such as the alpha7 nicotinic receptor (&#945;7-nAChR), phosphorylated Tau (p-Tau), and amyloid beta (A&#946;).

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

We present a method which can&#160;provide further insights into the early events underlying neurodegeneration and based on the established ex-vivo brain technique, combining the advantages of in vivo and in vitro experiments. Moreover, it represents a unique opportunity for direct comparison of treated and untreated group in the same anatomical plane.

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, ...

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model animal for endocrine research, Drosophila melanogaster, which has sophisticated genetic tools and genome information, is being increasingly used. This article describes a method for the calcium imaging of Drosophila brain explants. This method enables the detection of the direct signaling of a hormone to the brain. It is well known that many peptide hormones act through G-protein-coupled receptors (GPCRs), whose activation causes an increase in the intracellular Ca2+concentration. Neural activation also elevates intracellular Ca2+ levels, from both Ca2+ influx and the release of Ca2+ stored in the endoplasmic reticulum (ER). A calcium sensor, GCaMP, can monitor these Ca2+ changes. In this method, GCaMP is expressed in the neurons of interest, and the GCaMP-expressing larval brain is dissected and cultured ex vivo. The test peptide is then applied to the brain explant, and the fluorescent changes in GCaMP are detected using a spinning disc confocal microscope equipped with a CCD camera. Using this method, any water-soluble molecule can be tested, and various cellular events associated with neural activation can be imaged using the appropriate fluorescent indicators. Moreover, by modifying the imaging chamber, this method can be used to image other Drosophila organs or the organs of other animals.

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

This paper describes a protocol for ex vivo calcium imaging of the Drosophila brain. In this method, natural or synthetic compounds can be applied to the buffer to test their ability to activate particular neurons in the brain.

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model ...

Using Near-infrared Fluorescence and High-resolution Scanning to Measure Protein Expression in the Rodent Brain

Neuroscience is the study of how cells in the brain mediate various functions. Measuring protein expression in neurons and glia is critical for the study of neuroscience as cellular function is determined by the composition and activity of cellular proteins. In this article, we describe how immunocytochemistry can be combined with near-infrared high-resolution scanning to provide a semi-quantitative measure of protein expression in distinct brain regions. This technique can be used for single or double protein expression in the same brain region. Measuring proteins in this fashion can be used to obtain a relative change in protein expression with an experimental manipulation, molecular signature of learning and memory, activity in molecular pathways, and neural activity in multiple brain regions. Using the correct proteins and statistical analysis, functional connectivity among brain regions can be determined as well. Given the ease of implementing immunocytochemistry in a laboratory, using immunocytochemistry with near-infrared high-resolution scanning can expand the ability of the neuroscientist to examine neurobiological processes at a systems level.

Here, we present a protocol that uses near-infrared dyes in conjunction with immunohistochemistry and high-resolution scanning to assay proteins in brain regions.

Neuroscience is the study of how cells in the brain mediate various functions. Measuring protein expression in neurons and glia is critical for the study ...