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><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 ...

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Research

JoVE Journal

Neuroscience

3.2K Views.  Universidad de Atacama. Deze techniek is een aanpassing van veelgebruikte methoden voor het onderzoeken van monoamineafgifte, waardoor de endogene afgifte kan worden gemeten in plaats van vooraf geladen radioactief gelabelde monoaminen onder meerdere farmacologische omstandigheden tegelijk. De belangrijkste voordelen van deze techniek zijn dat het redelijk kosteneffectief is, meerdere farmacologische aandoeningen tegelijk kunnen worden onderzocht en we endogene monoamineafgifte kunnen bestuderen met het vermogen om ...

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 ...

Measurement of Glutamate Uptake using Radiolabeled L-[3H]-Glutamate in Acute Transverse Slices Obtained from Rodent Resected Hippocampus

Glutamate removal from the extracellular space by high-affinity Na+-dependent transporters is essential to ensure that the brain&#39;s intrinsic connectivity mechanisms work properly and homeostasis is maintained. The hippocampus is a unique brain structure that manages higher cognitive functions, and is the subject of several studies regarding neurologic diseases. The investigation of physiological and pathological mechanisms in rodent models can benefit from acute hippocampal slice (AHS) preparations. AHS has the advantage of providing reliable information on cell function since the cytoarchitecture and synaptic circuits are preserved. Although AHS preparations are commonly used in neurochemistry laboratories, it is possible to find some methodological differences in the literature. Considering that distinctive slice preparation protocols might change the hippocampal regions analyzed, this current protocol proposes a standard technique for obtaining transverse AHS from resected hippocampus. This simple-to-perform protocol may be used in mice and rats&#39; experimental models and allow several ex vivo approaches investigating neurochemical dynamics (in dorsal, intermediate and ventral hippocampus) in different backgrounds (e.g., transgenic manipulations) or after in vivo manipulations (e.g., pharmacological treatments or suitable rodent models to study clinical disorders). After dissecting the hippocampus from the rodent brain, transverse slices along the septo-temporal axis (300 &#181;m thick) were obtained. These AHS contain distinct parts of the hippocampus and were subjected to an individual neurochemical investigation (as an example: neurotransmitter transporters using their respective substrates). As the hippocampus presents a high density of excitatory synapses, and glutamate is the most important neurotransmitter in the brain, the glutamatergic system is an interesting target for in vivo observed phenomena. Thus, the current protocol provides detailed steps to explore glutamate uptake in ex vivo AHS using L-[3H]-Glutamate. Using this protocol to investigate hippocampal function may help to better understand the influence of glutamate metabolism on mechanisms of neuroprotection or neurotoxicity.

Measurement of Glutamate Uptake using Radiolabeled L-[3H]-Glutamate in Acute Transverse Slices Obtained from Rodent Resected Hippocampus

This article describes a reliable and simple way to obtain ex vivo acute hippocampal transverse slices from mice and rats using a tissue chopper. Slices obtained from resected hippocampi can be submitted for functional glutamate uptake analysis to investigate glutamatergic system homeostasis.

Glutamate removal from the extracellular space by high-affinity Na+-dependent transporters is essential to ensure that the brain&#39;s intrinsic ...

Biochemistry

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.

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.

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

Using Enzyme-based Biosensors to Measure Tonic and Phasic Glutamate in Alzheimer's Mouse Models

Neurotransmitter disruption is often a key component of diseases of the central nervous system (CNS), playing a role in the pathology underlying Alzheimer's disease, Parkinson's disease, depression, and anxiety. Traditionally, microdialysis has been the most common (lauded) technique to examine neurotransmitter changes that occur in these disorders. But because microdialysis has the ability to measure slow 1-20 minute changes across large areas of tissue, it has the disadvantage of invasiveness, potentially destroying intrinsic connections within the brain and a slow sampling capability. A relatively newer technique, the microelectrode array (MEA), has numerous advantages for measuring specific neurotransmitter changes within discrete brain regions as they occur, making for a spatially and temporally precise approach. In addition, using MEAs is minimally invasive, allowing for measurement of neurotransmitter alterations in vivo. In our laboratory, we have been specifically interested in changes in the neurotransmitter, glutamate, related to Alzheimer's disease pathology. As such, the method described here has been used to assess potential hippocampal disruptions in glutamate in a transgenic mouse model of Alzheimer's disease. Briefly, the method used involves coating a multi-site microelectrode with an enzyme very selective for the neurotransmitter of interest and using self-referencing sites to subtract out background noise and interferents. After plating and calibration, the MEA can be constructed with a micropipette and lowered into the brain region of interest using a stereotaxic device. Here, the method described involves anesthetizing rTg(TauP301L)4510 mice and using a stereotaxic device to precisely target sub-regions (DG, CA1, and CA3) of the hippocampus.

Here, we describe the setup, software navigation, and data analysis for a spatially and temporally precise method of measuring tonic and phasic extracellular glutamate changes in vivo using enzyme-linked microelectrode arrays (MEA).

Neurotransmitter disruption is often a key component of diseases of the central nervous system (CNS), playing a role in the pathology underlying ...

Assessment of Dopaminergic Homeostasis in Mice by Use of High-performance Liquid Chromatography Analysis and Synaptosomal Dopamine Uptake

Dopamine (DA) is a modulatory neurotransmitter controlling motor activity, reward processes and cognitive function. Impairment of dopaminergic (DAergic) neurotransmission is strongly associated with several central nervous system-associated diseases such as Parkinson&#39;s disease, attention-deficit-hyperactivity disorder and drug addiction1,2,3,4. Delineating disease mechanisms involving DA imbalance is critically dependent on animal models to mimic aspects of the diseases, and thus protocols that assess specific parts of the DA homeostasis are important to provide novel insights and possible therapeutic targets for these diseases.
Here, we present two useful experimental protocols that when combined provide a functional read-out of the DAergic system in mice. Biochemical and functional parameters on DA homeostasis are obtained through assessment of DA levels and dopamine transporter (DAT) functionality5. When investigating the DA system, the ability to reliably measure endogenous levels of DA from adult brain is essential. Therefore, we present how to perform high-performance liquid chromatography (HPLC) on brain tissue from mice to determine levels of DA. We perform the experiment on tissue from dorsal striatum (dStr) and nucleus accumbens (NAc), but the method is also suitable for other DA-innervated brain areas.
DAT is essential for reuptake of DA into the presynaptic terminal, thereby controlling the temporal and spatial activity of released DA. Knowing the levels and functionality of DAT in the striatum is of major importance when assessing DA homeostasis. Here, we provide a protocol that allows to simultaneously deduce information on surface levels and function using a synaptosomal6 DA uptake assay.
Current methods combined with standard immunoblotting protocols provide the researcher with relevant tools to characterize the DAergic system.

Synaptosomal dopamine uptake and high-performance liquid chromatography analysis represent experimental tools to investigate dopamine homeostasis in mice by assessing the function of the dopamine transporter and levels of dopamine in striatal tissue, respectively. Here we present protocols to measure dopamine tissue content and assess the functionality of the dopamine transporter.

Dopamine (DA) is a modulatory neurotransmitter controlling motor activity, reward processes and cognitive function. Impairment of dopaminergic (DAergic) ...

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.

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other ...

Consistent Induction and Recording of Seizure-Like Activity Using CMOS-HD-MEAs

High-Quality Seizure-Like Activity from Acute Brain Slices Using a Complementary Metal-Oxide-Semiconductor High-Density Microelectrode Array System

Complementary metal-oxide-semiconductor high-density microelectrode array (CMOS-HD-MEA) systems can record neurophysiological activity from cell cultures and ex vivo brain slices in unprecedented electrophysiological detail. CMOS-HD-MEAs were first optimized to record high-quality neuronal unit activity from cell cultures but have also been shown to produce quality data from acute retinal and cerebellar slices. Researchers have recently used CMOS-HD-MEAs to record local field potentials (LFPs) from acute, cortical rodent brain slices. One LFP of interest is seizure-like activity. While many users have produced brief, spontaneous epileptiform discharges using CMOS-HD-MEAs, few users reliably produce quality seizure-like activity. Many factors may contribute to this difficulty, including electrical noise, the inconsistent nature of producing seizure-like activity when using submerged recording chambers, and the limitation that 2D CMOS-MEA chips only record from the surface of the brain slice. The techniques detailed in this protocol should enable users to consistently induce and record high-quality seizure-like activity from acute brain slices with a CMOS-HD-MEA system. In addition, this protocol outlines the proper treatment of CMOS-HD-MEA chips, the management of solutions and brain slices during experimentation, and equipment maintenance.

Author Spotlight: Unraveling Seizure Dynamics and Novel Therapeutics for Status Epilepticus Using CMOS High-Density Microelectrode Array Systems

Here, we outline a protocol for using complementary metal-oxide-semiconductor high-density microelectrode array systems (CMOS-HD-MEAs)&#160;to record seizure-like activity from ex vivo brain slices.

Complementary metal-oxide-semiconductor high-density microelectrode array (CMOS-HD-MEA) systems can record neurophysiological activity from cell cultures ...

Isolation of the Brain Secretome from Ex Vivo Brain Slice Cultures

The brain secretome consists of proteins either actively secreted or shed from the cell surface by proteolytic cleavage in the extracellular matrix of the nervous system. These proteins include growth factor receptors and transmembrane proteins, among others, covering a broad spectrum of roles in the development and normal functioning of the central nervous system. The current procedure to extract the secretome from cerebrospinal fluid is complicated and time-consuming, and it is difficult to isolate these proteins from experimental animal brains. In this study, we present a novel protocol for isolating the brain secretome from mouse brain slice cultures. First, the brains were isolated, sliced, and cultured ex vivo. The culture medium was then filtered and concentrated for isolating proteins by centrifugation after a few days. Finally, the isolated proteins were resolved using sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and subsequently probed for purity characterization by western blot. This isolation procedure of the brain secretome from ex vivo brain slice cultures can be used to investigate the effects of the secretome on a variety of neurodevelopmental diseases, such as autism spectrum disorders.

Isolation of the Brain Secretome from Ex Vivo Brain Slice Cultures

The brain secretome plays a pivotal role in the development and normal functioning of the central nervous system. In this article, we provide a detailed protocol for isolation of the brain secretome from ex vivo brain slice cultures.

The brain secretome consists of proteins either actively secreted or shed from the cell surface by proteolytic cleavage in the extracellular matrix of the ...

Biology

A Plate-Based Assay to Measure Monoamine Release from Brain Slices on Test Drug Exposure

Source: Pino, J. A. et al. <a href="https://app.jove.com/v/62127">A Plate-Based Assay for the Measurement of Endogenous Monoamine Release in Acute Brain Slices.</a> J. Vis. Exp. (2021)
In this video, we demonstrate an assay that investigates monoamine release mechanisms from the neurons of metabolically active ex vivo rat brain slices, when treated with test drugs. The assay demonstrates that neurostimulant drugs induce monoamine release via the cellular monoamine transporters, whereas high KCl concentration induces synaptic vesicle exocytosis-mediated vesicular monoamine release.

Source: Pino, J. A. et al. A Plate-Based Assay for the Measurement of Endogenous Monoamine Release in Acute Brain Slices. J. Vis. Exp. (2021)
In this ...

JoVE Encyclopedia of Experiments

Biological Techniques

Assay Techniques

Biochemical assays

Measuring Dopamine Uptake in Mouse Brain Synaptosomes Using Radiolabeled Dopamine

Source: Jensen, K. L., et al., <a href="https://app.jove.com/v/56093">Assessment of Dopaminergic Homeostasis in Mice by Use of High-performance Liquid Chromatography Analysis and Synaptosomal Dopamine Uptake.</a> J. Vis. Exp. (2017).
This video demonstrates the assessment of dopamine uptake in synaptosomes harvested from mouse brains. It outlines steps for preparing synaptosomes for dopamine uptake via dopamine transporters, treating them with varying concentrations of radiolabeled dopamine, isolating the synaptosomes, and measuring radioactivity to quantify dopamine uptake.

Source: Jensen, K. L., et al., Assessment of Dopaminergic Homeostasis in Mice by Use of High-performance Liquid Chromatography Analysis and Synaptosomal ...

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

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