This work was supported by grants Fondecyt Initiation Fund N 11191049 to J.A.P. and NIH grant DA038598 to G.E.T.

All experiments, including animal handling and tissue collection, were carried out in accordance with the University of Florida and the City College of New York Institutional Animal Care and Use Committee (IACUC), following the approved protocol 201508873 (UF) and 1071 (CCNY). For reagents and buffer please refer to the Supplementary File.
1. Prepare acute rat brain slices
NOTE: In this experiment adult male rats (250-350 g) were used. However, this set up is functional for different developmental points, female rats, and other species. If using a smaller animal, such as mice, the experimenter may adjust to optimize the protocol by using a different number of brain slices or punches per condition. Dissection buffer will be referred to as Buffer 1; efflux buffer will be referred to as Buffer 2.
<ol>
	<li>Prepare Buffer 1 as mentioned in the Supplementary File. Saturate Buffer 1 with oxygen by bubbling with 95%/5% (O2/CO2) for 20 min on ice. Remove 50 mL of Buffer 1, and chill on ice in a small beaker or a Petri dish. This buffer is used to hold the acutely harvested whole brain.</li>
	<li>Anesthetize one or two adult rats (250-350 g) with 1%-2% isoflurane, decapitate them using a guillotine, and rapidly remove their brains. Immediately place the brain in ice-cold oxygenated Buffer 1 in the container from step 1.1. 
	NOTE: Ensure isoflurane and guillotine are used safely. Open isoflurane under a fume hood.</li>
	<li>Using a vibratome or compresstome, cut 300 &#181;m coronal brain sections from each region of interest (Figure 1). Bubbling Buffer 1 must be present while sections are being made. Using a stainless-steel spatula, carefully and immediately transfer brain slices into a new Petri dish filled with ice-cold oxygenated Buffer 1 (Figure 2).</li>
	<li>Further dissect brain slices (e.g., punches, cut out) by carefully moving the slices to glass slides (Figure 1G) using rat brain atlas57. For instance, identify the dorsal striatum based off its dark, striated structure, and identify the hippocampus based off its proximity to the cortex and unique spiral structure. The right and left hemispheres may be separated to use as control and experimental slices (Figures 2G - H). Here, the dorsal striatum was further dissected into 2 mm punches (Figure 1G).</li>
	<li>Using a plastic transfer pipette with the tip cut off, transfer slices or brain punches into small containers immersed in oxygenated ice-cold Buffer 1 with oxygen bubbling. These containers may be stainless steel mesh, or small Petri dishes filled with buffer (Figure 1H).</li>
</ol>
2. Ex-vivo endogenous monoamine release from brain slices or punches
NOTE: The device used for this section consists of a 48-well plate and a tissue holder made of six microcentrifuge filter units without the inset-filters connected to a carbogen line (Figure 2). To make the holder, use a sturdy plastic rod (e.g., from a cell scrapper) and super glue the microcentrifuge filter units without the inset-filters to it. Let it dry for 1-2 days. Time required for the endogenous monoamine release experiment and concentrations of amphetamine, fluoxetine, and cocaine are based on the current literature and previous protocols13,20,58.
<ol>
	<li>Tissue activation
	<ol>
		<li>Transfer the brain tissue from step 1.1.5 to each well of the efflux chamber and allow recovery for 30-50 min at 37 &#176;C on a slide warmer in 0.5-1 mL of oxygenated Buffer 2, with constant, gentle bubbling (Figure 2B1).</li>
		<li>During this incubation, dilute the drugs to the desired concentration for the experiment. All the drugs must be dissolved in Buffer 2, and concentrations are based off the current literature.</li>
	</ol>
	</li>
	<li>First incubation
	<ol>
		<li>Move the tissue holder with brain tissue to wells containing 500 &#181;L of oxygenated Buffer 2 and incubate for 20 min at 37 &#176;C. Ensure that minimal to no buffer is transported over by tapping the holder on the edge of the well until no excess buffer is in the holder.</li>
		<li>In experiments with pharmacological agents such as monoamine transporter inhibitors, incubate the tissue samples with the drugs diluted in oxygenated Buffer 2 (e.g., 10 &#181;M Fluoxetine, 40 &#181;M cocaine; see Figure 2B2). The final volume in each well will be 500 &#181;L.</li>
	</ol>
	</li>
	<li>Second incubation
	<ol>
		<li>Move the holder with the tissue to a new set of wells containing 500 &#181;L total Buffer 2 with or without the desired concentration of each drug. Ensure there is no excess buffer leftover. Each well represents an n = 1 for experimental conditions. Each experimental condition is performed in triplicate.</li>
		<li>One well includes an oxygenated Buffer 2, the next 10-30 &#181;M AMPH, and the final well includes 10-30 &#181;M AMPH plus monoamine transporter inhibitors. Each drug is dissolved in oxygenated Buffer 2.</li>
		<li>Incubate the tissue for 20 min at 37 &#176;C with 500 &#181;L of the drug condition. 
		&#8203;NOTE: Additional wells may include an oxygenated high K+ Buffer 2 with or without the monoamine transporter inhibitors. Dissolve each drug in the oxygenated Buffer 2 (500 &#181;L).</li>
		<li>During this second incubation of 20 min, collect the solution from the wells from the first incubation in step 2.2.1 and transfer to microcentrifuge tubes containing 50 &#181;L of 1 N perchloric acid or phosphoric acid (dependent on the type of HPLC, final concentration 0.1 N). The final volume of the sample will be 550 &#181;L. Maintain microcentrifuge tubes on ice, and label the tubes #1.</li>
		<li>After the second incubation of 20 min, move the tissue holder with brain sections or punches to an empty well and maintain the plate on ice. Transfer the supernatant to microcentrifuge tubes containing 50 &#181;L of 1 N perchloric acid or phosphoric acid. The final volume of the sample will be 550 &#181;L. Maintain microcentrifuge tubes on ice, and label the tubes #2.</li>
		<li>Add 1 mL of ice-cold Buffer 1 to each well containing tissue. Collect all of the tissue using small tweezers, and transfer to clean microcentrifuge tubes.</li>
		<li>Maintain tubes with brain tissue at -80 &#176;C. Discard the 1 mL of Buffer 1 (Figure 2B4).</li>
		<li>Filter solutions obtained from each incubation using microcentrifuge filter tubes (0.22 &#181;m) at 2,500 x g for 2min. Use the filtrate to determine monoamine content using HPLC with electrochemical detection (Figure 2B5).</li>
	</ol>
	</li>
</ol>
3. Tissue viability
<ol>
	<li>MTT assay 
	&#8203;NOTE: A significant concern regarding this experimental setup is tissue viability as the tissue may be used for up to several hours59. An MTT assay60,61 is used to determine tissue viability by the end of the experimentation. This assay is based on the conversion of the yellow tetrazolium salt MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) to purple formazan crystals by viable cells with adequate metabolism.
	<ol>
		<li>Post experiment maintain a separate group of tissue samples and separate them into two groups.</li>
		<li>Incubate one group for 20 min at 37 &#176;C in Triton X-100 (1%) dissolved in Buffer 2 as a control. Triton X-100 treatment results in cell death. Maintain the second group in Buffer 2, and do not incubate in Triton X-100 (tissue viability control).</li>
		<li>Add MTT (stock solution 5 mg/mL in PBS, pH 7.4) to both groups in the oxygenated Buffer 2 to a final concentration of 0.5 mg/mL.</li>
		<li>Incubate the tissue samples for 20 min at 37 &#176;C, wash with PBS, and transfer into microcentrifuge tubes containing 250 &#181;L of a mixture of SDS (10%, w/v), DMF (25%, v/v), and water to dissolve the formazan crystals.</li>
		<li>Incubate the samples for 24 h.</li>
		<li>Centrifuge the tubes at 10,000 x g for 10 min and measure the absorbance of the supernatant (200 &#181;L) at 562 nm and 690 nm using a microplate reader. Tissue viability is calculated as follows: (A562-A690)/tissue weight.</li>
	</ol>
	</li>
</ol>
4. HPLC analysis of monoamines
<ol>
	<li>Quantify monoamine release from each experimental condition using HPLC-ECD according to previous protocols13,44, using a reverse phase column.
	<ol>
		<li>Prepare the mobile phase required for detection. This consists of 100 mM phosphoric acid, 100 mM citric acid, 0.1 mM EDTA-Na2, 600 mg/L octanesulfonic acid, 8% v/v acetonitrile (final pH 6.0). The composition of the mobile phase is dependent on type of HPLC and column used.</li>
		<li>Set the potential of the electrochemical detector (2 mm glassy carbon electrode,) to 0.46 V, and set the flow rate to 0.05 mL/min.</li>
		<li>Load 5 &#181;L of each sample, including neurotransmitter standards into the HPLC for autoinjection and detection. The amount of each sample added depends on the type of HPLC used.</li>
		<li>Once the HPLC has completed the run, use the given HPLC analysis software to acquire and analyze the chromatograph data.</li>
		<li>Analyze monoamine content using a standard curve composed of each monoamine (Dopamine: DA, Norepinephrine: NE, and Serotonin: 5-HT; Figure 2C). Use the resulting chromatograms to obtain the area under the curve (AUC) based on the manufacturer&#39;s guidelines.</li>
	</ol>
	</li>
</ol>
5. Preparing tissue lysates for protein quantification
<ol>
	<li>Protein assay
	<ol>
		<li>Add ice-cold lysis buffer plus protease inhibitors (0.1 g/1 mL) to each microcentrifuge tube containing brain sections/punches and homogenize using a pestle homogenizer. The microcentrifuge tubes must be maintained on ice while homogenizing to prevent protein degradation.</li>
		<li>Incubate tissue homogenates for 1 h at 4 &#176;C with light rotation.</li>
		<li>Centrifuge tissue homogenates at 16,000 x g for 15 min at 4 &#176;C and recover the supernatant.</li>
		<li>Determine protein concentration in the supernatants, with bovine serum albumin (BSA) as a standard.</li>
		<li>Normalize the monoamine content in each brain sample to the total content of protein (&#181;g) measured in 250 &#181;L of brain tissue lysed. Use the below formula to determine the nmol monoamine/g protein. df = dilution factor.</li>
	</ol>
	</li>
</ol>
<img alt="Equation 1" src="/files/ftp_upload/62127/62127eq01.jpg" />
6. Statistical analysis
<ol>
	<li>Analyze monoamine release (nmol/g) using one-way ANOVA followed by Sidak&#39;s multiple comparisons test for post-hoc comparisons.</li>
	<li>Analyze tissue viability using an Unpaired Student&#39;s t-test for independent groups (Control vs. 1% Triton X-100).</li>
	<li>For all statistical analyses, set the alpha level to &#8804; 0.05.</li>
</ol>

The authors have no disclosures.

Monoamine release measurements have been performed for years in a number of systems such as heterologous cells, neuronal cultures, brain synaptosomes, ex vivo acute brain slices, and whole animals13,20,41,42,58,64,65,66,67...

A plethora of neurological and psychiatric diseases are associated with dysregulation or insufficient maintenance of monoamine neurotransmitter (dopamine [DA], serotonin [5-HT], norepinephrine [NE]) homeostasis1,2,3. These conditions include, but are not limited to, depression1,2, schizophrenia2, anxiety2, addiction4, menopause5,6,7, pain8, and Parkinson&#39;s disease3. For instance, several rat models of menopause have shown that the dysregulation or reduction of monoamines within the hippocampus, prefrontal cortex, and striatum may be associated with both depression and cognitive decline, which is seen in women experiencing menopause. The dysregulation of monoamines in these models have been extensively examined using HPLC-ECD, although the studies did not discriminate between measured neurotransmitter content versus neurotransmitter release5,6,7. Monoamines are classically released into the extracellular space through Ca2+-dependent vesicular release9, and are recycled back through their respective plasma membrane re-uptake system (dopamine transporter, DAT; serotonin transporter, SERT; norepinephrine transporter, NET)10,11. Conversely, data suggests that these transporters are able to release or efflux monoamines, since drugs of abuse such as amphetamine (AMPH) and 3,4-Methylenedioxymethamphetamine (MDMA) are known to release DA and 5-HT, respectively through their transporter systems12,13,14,15,16,17. Thus, a proper mechanistic understanding of monoamine release dynamics is crucial for developing specific and targeted pharmacotherapies.
A wide range of techniques have been employed to study monoamine release such as Fast Scan Cyclic Voltammetry (FSCV)18, in vivo&#160;microdialysis13, imaging19, preincubation with radiolabeled monoamines20, optogenetics, and more recently, genetically encoded fluorescent sensors and photometry21,22. FSCV and in vivo microdialysis are the primary techniques used for studying monoamine release. FSCV is used to study the stimulated exocytotic release of, primarily, DA in acute brain slices and in vivo23. Because FSCV uses electrodes to stimulate or evoke release, the primary source of neurotransmitter release is Ca2+-dependent vesicular release18,24,25,26,27,28,29,30,31. In vivo&#160;microdialysis coupled with HPLC measures changes in extracellular neurotransmitter levels using a probe placed in a brain area of interest13,32. Similar to FSCV, a major limitation to in vivo&#160;microdialysis is the difficulty in determining the source of neurotransmitter release: Ca2+ dependent vesicular release or transporter dependent. Noteworthy, both methods allow for the direct measurement of monoamine release. Through the recent advancement of optogenetics, research demonstrates detection of 5-HT and DA release in a short timespan with exquisite cell-type specificity21,22. However, these strategies require complex and costly techniques and equipment, and indirectly measure monoamine release, specifically through monoamine binding to receptors. Further, radiolabeled monoamines are also used for studying monoamine dynamics. Radiolabeled monoamines may be preloaded into various model systems such as heterologous cells overexpressing each monoamine transporter20,33,34,35,36,37,38,39,40, primary neurons20, synaptosomes33,39,41,42, and acute brain slices43,44. However, radioactivity poses potential harm to the experimenter, and the tritium-labeled analytes may not faithfully recapitulate endogenous monoamine dynamics45,46. Superfusion systems combined with off-line detection methods such as HPLC-ECD have allowed for the detection of monoamines from multiple tissue sources. Here, this protocol provides as an optimized and low-cost, simple, and precise method using acute brain slices to directly measure endogenous basal and stimulated monoamine release.
Acute brain slices allow for testing mechanistic hypotheses, primarily as they preserve the in vivo anatomical microenvironment, and maintain intact synapses47,48,49,50,51,52. In a few studies, acute brain slices or chopped brain tissue have been used in conjunction with a superfusion technique using KCl to stimulate Ca2+ mediated release53,54,55,56. Superfusion systems have been critical to advance the field&#39;s understanding of neurotransmitter release mechanisms, including monoamines. However, these systems are relatively expensive, and the number of chambers available for tissue analysis ranges from 4-12. In comparison, the method presented here is inexpensive, allows the measurement of 48 tissue samples, and may be refined to use up to 96 tissue samples. Each well within the 48-well plate contains tissue holders that use filters to separate the released product from the tissue, and released monoamines are then collected and analyzed by HPLC-ECD. Importantly, this method allows for the simultaneous measurement of 5-HT, DA, and NE release from different brain areas such as the prefrontal cortex, the hippocampus, and the dorsal striatum after treatment with pharmacological agents that modulate monoamine release. Thus, the experimenter can answer multiple questions using an inexpensive multi-well system that increases the number of samples tested and thereby reducing the number of animals used.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>48 Well plate</td>
			<td>NA</td>
			<td>NA</td>
			<td>Assay</td>
		</tr>
		<tr>
			<td>Acetonitrile</td>
			<td>Fischer Scientific</td>
			<td>A998-1</td>
			<td>Mobile Phase</td>
		</tr>
		<tr>
			<td>Calcium Chloride Ahydrous</td>
			<td>Sigma Aldrich</td>
			<td>C1016</td>
			<td>Modified Artifical Cerebrospinal Fluid OR Efflux Buffer</td>
		</tr>
		<tr>
			<td>Clarity Software</td>
			<td>Anetc</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Citric Acid</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>Mobile Phase</td>
		</tr>
		<tr>
			<td>D-(+)-Glucose</td>
			<td>Sigma</td>
			<td>1002608421</td>
			<td>Dissection Buffer</td>
		</tr>
		<tr>
			<td>DMF</td>
			<td>Sigma Aldrich</td>
			<td>D4551</td>
			<td>MTT Assay</td>
		</tr>
		<tr>
			<td>EDTA-Na2</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>Mobile Phase</td>
		</tr>
		<tr>
			<td>GraphPad Software</td>
			<td>Graphpad Software, Inc</td>
			<td></td>
			<td>Statistical Analysis</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Sigma Aldrich</td>
			<td>G5516</td>
			<td>Lysis buffer</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma Aldrich</td>
			<td>H3375</td>
			<td>Lysis buffer</td>
		</tr>
		<tr>
			<td>HPLC, Decade Amperometric</td>
			<td>Anetc</td>
			<td></td>
			<td>HPLC, LC-EC system</td>
		</tr>
		<tr>
			<td>HPLC</td>
			<td>Amuza</td>
			<td></td>
			<td>HPLC HTEC-510.</td>
		</tr>
		<tr>
			<td>L-Asrobic Acid</td>
			<td>Sigma Aldrich</td>
			<td>A5960</td>
			<td>Dissection Buffer</td>
		</tr>
		<tr>
			<td>Magnesium Sulfate</td>
			<td>Sigma</td>
			<td>7487-88-9</td>
			<td>KH Buffer</td>
		</tr>
		<tr>
			<td>Microcentrifuge Filter Units UltraFree</td>
			<td>Millipore</td>
			<td>C7554</td>
			<td>Assay - 6 to fit in 48 well plate</td>
		</tr>
		<tr>
			<td>MTT</td>
			<td>Thermo Fisher</td>
			<td>M6494</td>
			<td>MTT Assay</td>
		</tr>
		<tr>
			<td>Nanosep</td>
			<td>VWR</td>
			<td>29300-606</td>
			<td>Assay; protein assay</td>
		</tr>
		<tr>
			<td>Octanesulfonic acid</td>
			<td>Sigma Aldrich</td>
			<td>V800010</td>
			<td>Mobile Phase</td>
		</tr>
		<tr>
			<td>Pargyline Clorohydrate</td>
			<td>Sigma Aldrich</td>
			<td>P8013</td>
			<td>Modified Artifical Cerebrospinal Fluid OR Efflux Buffer</td>
		</tr>
		<tr>
			<td>Phosphoric Acid</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>Mobile Phase</td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>Sigma</td>
			<td>12636</td>
			<td>KH Buffer</td>
		</tr>
		<tr>
			<td>Potassium Phosphate Monobasic</td>
			<td>Sigma</td>
			<td>1001655559</td>
			<td>KH Buffer</td>
		</tr>
		<tr>
			<td>Precisonary VF-21-0Z</td>
			<td>Precissonary</td>
			<td></td>
			<td>Compresstome</td>
		</tr>
		<tr>
			<td>Protease Inhibitor Cocktail</td>
			<td>Sigma Aldrich</td>
			<td>P2714</td>
			<td>Lysis buffer.</td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate</td>
			<td>Sigma</td>
			<td>S5761</td>
			<td>Dissection Buffer</td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate</td>
			<td>Sigma Aldrich</td>
			<td>S5761</td>
			<td>Dissection Buffer</td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Sigma</td>
			<td>S3014</td>
			<td>KH Buffer</td>
		</tr>
		<tr>
			<td>Sodium Dodecyl Sulfate</td>
			<td>Sigma Aldrich</td>
			<td>L3771</td>
			<td>Lysis buffer</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma Aldrich</td>
			<td>T8787</td>
			<td>MTT Assay / Lysis buffer</td>
		</tr>
	</tbody>
</table>

a plate-based assay for the measurement of endogenous monoamine release in acute brain slices

Monoamine neurotransmitters are associated with numerous neurologic and psychiatric ailments. Animal models of such conditions have shown alterations in monoamine neurotransmitter release and uptake dynamics. Technically complex methods such as electrophysiology, Fast Scan Cyclic Voltammetry (FSCV), imaging, in vivo&#160;microdialysis, optogenetics, or use of radioactivity are required to study monoamine function. The method presented here is an optimized two-step approach for detecting monoamine release in acute brain slices using a 48-well plate containing tissue holders for examining monoamine release, and high-performance liquid chromatography coupled with electrochemical detection (HPLC-ECD) for monoamine release measurement. Briefly, rat brain sections containing regions of interest, including prefrontal cortex, hippocampus, and dorsal striatum were obtained using a tissue slicer or vibratome. These regions of interest were dissected from the whole brain and incubated in an oxygenated physiological buffer. Viability was examined throughout the experimental time course, by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. The acutely dissected brain regions were incubated in varying drug conditions that are known to induce monoamine release through the transporter (amphetamine) or through the activation of exocytotic vesicular release (KCl). After incubation, the released products in the supernatant were collected and analyzed through an HPLC-ECD system. Here, basal monoamine release is detected by HPLC from acute brain slices. This data supports previous in vivo and in vitro results showing that AMPH and KCl induce monoamine release. This method is particularly useful for studying mechanisms associated with monoamine transporter-dependent release and provides an opportunity to screen compounds affecting monoamine release in a rapid and low-cost manner.

This technique describes the use of brain slices to measure the release of endogenous monoamines using HPLC with electrochemical detection based in a 48-well plate with an internal tissue holder. Experimental set up is depicted in Figure 1 and Figure 2. Initially, to ensure tissue viability by the end of the experimentation, an MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide, a tetrazole) assay was performed. Af...

Watch this Scientific Journal Video about A Plate-Based Assay for the Measurement of Endogenous Monoamine Release in Acute Brain Slices at JoVE.com

A Plate-Based Assay for the Measurement of Endogenous Monoamine Release in Acute Brain Slices

This method introduces a simple technique for the detection of endogenous monoamine release using acute brain slices. The setup uses a 48-well plate containing a tissue holder for monoamine release. Released monoamine is analyzed by HPLC coupled with electrochemical detection. Additionally, this technique provides a screening method for drug discovery.

Monoamine neurotransmitters are associated with numerous neurologic and psychiatric ailments. Animal models of such conditions have shown alterations in ...

a-plate-based-assay-for-measurement-endogenous-monoamine-release

Research

JoVE Journal

Neuroscience

3.2K Views.  Universidad de Atacama. This method introduces a simple technique for the detection of endogenous monoamine release using acute brain slices. The setup uses a 48-well plate containing a tissue holder for monoamine release. Released monoamine is analyzed by HPLC coupled with electrochemical detection. Additionally, this technique provides a screening method for drug discovery.

Video: A Plate-Based Assay for the Measurement of Endogenous Monoamine Release in Acute Brain Slices

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Preparation of Acute Subventricular Zone Slices for Calcium Imaging

The subventricular zone (SVZ) is one of the two neurogenic zones in the postnatal brain. The SVZ contains densely packed cells, including neural progenitor cells with astrocytic features (called SVZ astrocytes), neuroblasts, and intermediate progenitor cells. Neuroblasts born in the SVZ tangentially migrate a great distance to the olfactory bulb, where they differentiate into interneurons. Intercellular signaling through adhesion molecules and diffusible signals play important roles in controlling neurogenesis. Many of these signals trigger intercellular calcium activity that transmits information inside and between cells. Calcium activity is thus reflective of the activity of extracellular signals and is an optimal way to understand functional intercellular signaling among SVZ cells.
Calcium activity has been studied in many other regions and cell types, including mature astrocytes and neurons. However, the traditional method to load cells with calcium indicator dye (i.e. bath loading) was not efficient at loading all SVZ cell types. Indeed, the cellular density in the SVZ precludes dye diffusion inside the tissue. In addition, preparing sagittal slices will better preserve the three-dimensional arrangement of SVZ cells, particularly the stream of neuroblast migration on the rostral-caudal axis.
Here, we describe methods to prepare sagittal sections containing the SVZ, the loading of SVZ cells with calcium indicator dye, and the acquisition of calcium activity with time-lapse movies. We used Fluo-4 AM dye for loading SVZ astrocytes using pressure application inside the tissue. Calcium activity was recorded using a scanning confocal microscope allowing a precise resolution for distinguishing individual cells. Our approach is applicable to other neurogenic zones including the adult hippocampal subgranular zone and embryonic neurogenic zones. In addition, other types of dyes can be applied using the described method.

A method to load subventricular zone (SVZ) cells with calcium indicator dyes for recording calcium activity is described. The postnatal SVZ contains tightly packed cells including neural progenitor cells and neuroblasts. Rather than using bath loading we injected the dye by pressure inside the tissue allowing better dye diffusion.

The subventricular zone (SVZ) is one of the two neurogenic zones in the postnatal brain. The SVZ contains densely packed cells, including neural ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

In-depth Physiological Analysis of Defined Cell Populations in Acute Tissue Slices of the Mouse Vomeronasal Organ

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory neurons (VSNs) express a specific type of G protein-coupled receptor (GPCR) from at least three different chemoreceptor gene families allowing sensitive and specific detection of chemosensory cues. These families comprise the V1r and V2r gene families as well as the formyl peptide receptor (FPR)-related sequence (Fpr-rs) family of putative chemoreceptor genes. In order to understand the physiology of vomeronasal receptor-ligand interactions and downstream signaling, it is essential to identify the biophysical properties inherent to each specific class of VSNs.
The physiological approach described here allows identification and in-depth analysis of a defined population of sensory neurons using a transgenic mouse line (Fpr-rs3-i-Venus). The use of this protocol, however, is not restricted to this specific line and thus can easily be extended to other genetically modified lines or wild type animals.

Here, we describe a physiological approach that allows identification and in-depth analysis of a defined population of sensory neurons in acute coronal tissue slices of the mouse vomeronasal organ using whole-cell patch-clamp recordings.

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory ...

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 ...

Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a challenging and low-throughput technique due to its complexity and high reliance on user operation and control. This manuscript demonstrates an image-guided automatic patch clamp system for in vitro whole-cell patch clamp experiments in acute brain slices. Our system implements a computer vision-based algorithm to detect fluorescently labeled cells and to target them for fully automatic patching using a micromanipulator and internal pipette pressure control. The entire process is highly automated, with minimal requirements for human intervention. Real-time experimental information, including electrical resistance and internal pipette pressure, are documented electronically for future analysis and for optimization to different cell types. Although our system is described in the context of acute brain slice recordings, it can also be applied to the automated image-guided patch clamp of dissociated neurons, organotypic slice cultures, and other non-neuronal cell types.

This protocol describes how to conduct automatic image-guided patch-clamp experiments using a system recently developed for standard in vitro electrophysiology equipment.

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a ...

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from parallel fibers (PF) to Purkinje cells (PC) is considered the basis for motor learning, and deficiencies of both LTD and motor learning are observed in various gene-manipulated animals. Common motor learning sets, such as adaptation of the optokinetic reflex (OKR), the vestibular-ocular reflex (VOR), and rotarod test were used for evaluation of motor learning ability. However, results obtained from the GluA2-carboxy terminus modified knock-in mice demonstrated normal adaptation of the VOR and the OKR, despite lacking PF-LTD. In that report, induction of LTD was only attempted using one type of stimulation protocol at room temperature. Thus, conditions to induce cerebellar LTD were explored in the same knock-in mutants using various protocols at near physiological temperature. Finally, we found stimulation protocols, by which LTD could be induced in these gene-manipulated mice. In this study, a set of protocols are proposed to evaluate LTD-induction, which will more accurately allow examination of the causal relationship between LTD and motor learning. In conclusion, experimental conditions are crucial when evaluating LTD in gene-manipulated mice.

In some gene-manipulated animals, using a single protocol may fail to induce LTD in cerebellar Purkinje cells, and there may be a discrepancy between LTD and motor learning. Multiple protocols are necessary to assess LTD-induction in gene-manipulated animals. Standard protocols are shown.

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from ...

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.