We thank Wangchen Tsering and Pamela Walter for their critical reading and editing of this manuscript. This work was supported by the National Institute of Neurological Disorders and Stroke (NINDS) (NS054773 to C.J. L. and NS098393 to H.Z.) and the Department of Neuroscience at Thomas Jefferson University (Startup Funds to H.Z.).

All the procedures including animal work were conducted according to national and international guidelines and were approved by the Animal Care and Use Committee of Thomas Jefferson University. Male FVB/NTac mice at the age of 3 to 14 months were used. The following steps were performed in a non-sterile setting, but caution should be taken to keep everything as clean as possible.
NOTE: The method presented here was established and used in the research reported by Zhi et al.24. The experiments described here used a Seahorse XF24 extracellular flux analyzer (see Table of Materials). These methods can be adapted for the XFe24 analyzer, and some results were confirmed using this analyzer.
1. Hydrate cartridge sensors (1 day before the assay)
<ol>
	<li>Open the extracellular flux assay kit (see Table of Materials) and remove both the sensor cartridge (green) and utility plate (clear; Figure 1A). Place the sensor cartridge aside (sensors up) and do not touch the sensors.</li>
	<li>Add 600 &#181;L of calibrant solution (pH 7.4) to each well of the utility plate. Place the sensor cartridge on top of the utility plate and submerge the sensors in the calibrant solution. Make sure the triangular notch of the utility and sensor cartridge plate are correctly aligned (Figure 1B).</li>
	<li>Seal the extracellular flux assay kit with sealing film to prevent evaporation of the calibrant solution, and then place it in a 37 &#176;C incubator not supplemented with CO2 or oxygen overnight.</li>
</ol>
2. Prepare the tissue plate (on the day of the assay)
<ol>
	<li>Open the islet capture microplate and take out the islet plate for tissue sitting.</li>
	<li>Warm an appropriate volume (625 &#181;L per well) of pre-oxygenated modified artificial cerebrospinal fluid (ACSF, composition 140 mM NaCl, 2.5 mM KCl, 2.4 mM CaCl2, 1.3 mM MgSO4, 0.3 mM KH2PO4, 25 mM glucose, 10 mM HEPES, pH 7.2) buffer to 37 &#176;C in a 50 mL tube. Then add BSA to a final concentration of 4 mg/mL to prepare the respiration buffer. In general, 50 mL is enough for one plate.</li>
	<li>Add 625 &#181;L of respiration buffer (pre-oxygenated ACSF buffer with 25 mM glucose and 4 mg/mL BSA) to each well of the islet plate carefully and avoid shaking the plate. Ensure no residual drops are on top of the sensor cartridge and no air bubbles are present in the buffer of each well.</li>
</ol>
3. Acute striatal slice preparation and excised slice placement
<ol>
	<li>Decapitate the mouse after cervical dislocation and immediately dissect the brain in 10 mL of ice-cold pre-oxygenated cutting solution (125 mM NaCl, 2.5 mM KCl, 26 mM NaHCO3, 3.7 mM MgSO4, 0.3 mM KH2PO4, 10 mM glucose, pH 7.4).</li>
	<li>Section coronal striatal slices with a vibratome following the manufacturer&#39;s instruction (see Table of Materials) at a thickness of 150 &#181;m in ice-cold, pre-oxygenated cutting solution.</li>
	<li>Recover the slices in 50 mL of oxygenated ACSF (125 mM NaCl, 2.5 mM KCl, 26 mM NaHCO3, 2.4 mM CaCl2, 1.3 mM MgSO4, 0.3 mM KH2PO4, 10 mM glucose, 2 mM HEPES, pH 7.4) and keep in solution for up to 30 min at room temperature (RT).</li>
	<li>After recovery, transfer the slices to a 35 mm x 10 mm Petri dish with 5 mL of respiration buffer.</li>
	<li>Use a stainless-steel biopsy punch (e.g., 1.5 mm diameter) to create a circular piece of tissue in the desired area of the sliced brain. Keep the slice in the buffer while gently pressing down with the punch. Make sure the punch is pushed down hard enough to cut the tissue. Remove the rest of the tissue, lift the punch away, and remove the circular piece of tissue into the buffer.</li>
	<li>Cut the very end of a 1 mL pipette tip to make a hole with a 1.5-2.0 mm diameter and use it to hold and transfer the punched slice to the top of the capture screen. Suction one piece of punched brain tissue and carefully place the tissue onto the mesh side of the capture screen (Figure 2A). The capture screen will be a circular piece of plastic with the mesh attached to one side. 
	NOTE: The capture screens are included in the microplate pack (see Table of Materials) and are similar to the one used in Schuh et al.23.</li>
	<li>Gently use a paper tissue to dry the capture screen briefly. With the moisture removed, the tissue becomes sticky, which allows the slice to attach to the center of the mesh. Do not dry the slice too much as, otherwise, it will be damaged.</li>
	<li>Hold the capture screen slice side down with tweezers and place it into one of the wells of the incubating islet plate (Figure 2B). Be careful not to drop the slice; if so, take the screen out and place a new slice on it. 
	NOTE: Do not place brain slices into the two background correction wells (A1 and D6) in the islet plate.</li>
	<li>Incubate the islet plate at 37 &#176;C in an incubator for at least 30 min to allow temperature and pH equilibration before running the assay.</li>
</ol>
4. Loading of cartridges with desired compounds
<ol>
	<li>Dilute the desired compounds in modified ACSF&#160;(37 &#176;C) to the final stock concentration of 10x, 11x, 12x, and 13x of the working concentration for ports A to D, respectively, and adjust to pH 7.4. The test will administer the compounds in order from port A to D. For this experiment, the following solutions were used, with their respective working concentration mentioned before them: 10 mM pyruvate (port A), 20 &#181;M oligomycin (port B), 10 &#181;M or other concentrations of FCCP (port C), and 20 &#181;M antimycin A (port D).</li>
	<li>Gently preload 75 &#181;L of the diluted compounds into the appropriate injection ports of the sensor cartridge. Place the tips halfway into the injection ports at a 45&#176; angle with the tip against the wall of the injection port. Tips should not be inserted completely to the bottom of the injection ports as this may cause compound leakage through the port.</li>
	<li>Withdraw the tips from the ports carefully to avoid creating air bubbles. Do not tap any portion of the cartridge to avoid alleviating air bubbles.</li>
	<li>Visually inspect the injection ports for even loading. Ensure all liquid is in the port and no residual drops are present on top of the cartridge.</li>
	<li>Place the sensor cartridge onto the utility plate and put it into an incubator (non-CO2) for 30 min to allow it to heat up to 37 &#176;C again. Handle carefully by only holding onto the utility plate. Move as little as possible.</li>
</ol>
5. Calibration and performing the assay
<ol>
	<li>Load the assay template in the software. Press the green START button.</li>
	<li>Make sure to load the correct protocol. The protocol contains a combination of 3 min mix, 3 min wait, and 2 min measure sequences (these three steps form one measurement). Change the distance of the probe head from 26,600 to 27,800 in the hardware settings under instrument settings25. There is no need to change the probe head for the XFe24&#160;analyzer. Press START.</li>
	<li>Load the sensor cartridge on the utility plate into the instrument tray. The notch goes in the front, left corner. Ensure that the plate sits correctly and is flat. Load the drug-filled sensor cartridge into the analyzer for calibration.</li>
	<li>Follow the instructions on the screen in order to calibrate and equilibrate the sensors. This takes around 30 min.</li>
	<li>Once the calibration step is done, remove the calibration plate and replace it with the islet plate (containing the mesh and tissue slices).</li>
	<li>Measure the oxygen consumption in each well of the plate using the assay protocols. The protocol contains a combination of 3 min mix, 3 min wait, and 2 min measure sequences (these three steps form one measurement), at which point the OCR is calculated.</li>
	<li>For the entire experiment, include 4x OCR measurements to create a baseline, followed by the injection of port A (pyruvate) with 4x measurements, then the injection of port B (oligomycin) with 8x measurements, then the injection of port C (FCCP) with 5x measurements, and, finally, the injection of port D (antimycin A) with 6x measurements. 
	NOTE: This number of measurements after each compound injection is determined depending on when the measurement reaches the plateau.</li>
	<li>Analyze the OCR measurement data and the coupling efficiency data.</li>
</ol>

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The authors have nothing to disclose.

The method we developed allowed an XF analyzer to be used for measuring OCR in striatal slices from adult mice over a time span of 4 h. This method provides a new way to measure cellular bioenergetics in punches excised from anatomically defined brain structures. Since the tissue samples being analyzed are rather small, the metabolic parameters of specific brain areas involved in a disease can be investigated. In addition, using acute slices more closely mimics the physiological cellular environment, which cannot be achi...

Mitochondrial dysfunction has been implicated in several neurological diseases, including Parkinson&#39;s disease (PD), Huntington&#39;s disease, and Alzheimer&#39;s disease1,2,3. PD models such as PINK1 knockout (KO) mice and rats display impaired mitochondrial function4,5,6,7,8,9,10,11. Mitochondria isolated from the striatum (STR) or whole-brain of aged PINK1 KO mouse exhibit defects in complex I7,10,12,13. Directly measuring the oxygen consumption rate (OCR) is one of the most common methods to evaluate mitochondrial function since OCR is coupled with ATP production, the principal function of mitochondria14. Therefore, measuring OCR in disease models or patient-derived samples/tissue can help investigate how mitochondrial dysfunction leads to disease.
Currently, there are several ways to measure mitochondrial OCR, including the Clark electrode and other O2 electrodes, O2 fluorescent dye, and the extracellular flux analyzer15,16,17,18,19. As an advantage, O2 electrode-based methods allow various substrates to be easily added. However, they are insufficient for simultaneously measuring several samples. Compared to traditional O2 electrode-based methods, the extracellular flux analyzer, a commonly used tool for OCR in cell cultures or purified mitochondria, offers improved throughput15,18,20. Nevertheless, all these methods are usually applied to measure OCR in isolated mitochondria or cell cultures6,16,17,19,20,21. The isolation of mitochondria causes inadvertent damage, and extracted mitochondria or cell cultures are less physiologically relevant than intact brain slices22. Even when microelectrodes are used in slices, they are less sensitive and more difficult to operate than in cultured cells23.
To meet these challenges, we developed a method using the XF24 extracellular flux analyzer, which allows for the analysis of multiple metabolic parameters from acute striatal brain slices of mice24. This technique provides continuous direct quantification of mitochondrial respiration via the OCR. In brief, small sections of striatal brain slices are placed into wells of the islet plate, and the analyzer uses oxygen and proton fluorescent-based biosensors to measure the OCR and extracellular acidification rate, respectively17,21,25.
One of the unique features of the analyzer is the four injection wells, which allow the continued measurement of OCR while sequentially injecting up to four compounds or reagents; this enables the measurement of several cellular respiration parameters, such as basal mitochondrial OCR, ATP-linked OCR, and maximal mitochondrial OCR. The compounds injected during the measurements for the protocol shown here were working concentrations of 10 mM pyruvate in the first solution well (port A), 20 &#181;M oligomycin in the second solution well (port B), 10 &#181;M carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP) in the third well (port C), and 20 &#181;M antimycin A in the fourth well (port D), based on Fried et al.25. It must be noted that these concentrations were working concentrations, and stock solutions of 10x, 11x, 12x, and 13x were injected into the solution ports A through D, respectively. The purpose of using each solution was as follows: 1) Pyruvate was necessary since, without it, the addition of FCCP would have a decreased OCR response caused by a limitation of available substrates; 2) Oligomycin inhibits ATP synthase and allows the measurement of ATP linked respiration; 3) FCCP uncouples oxidation from phosphorylation and allows the measurement of maximal mitochondrial capacity; 4) Antimycin A inhibits complex III in the electron transport chain and, therefore, allows the measurement of OCR not linked to the mitochondria.
The concentration of oligomycin used was determined based on the following reasons: 1) The recommended dose of oligomycin for most cell types (isolated mitochondria or cell cultures) is 1.5 &#181;M. From experience, usually 3x-10x of the dissociated cells dose is used for the slice experiments since there might be a gradient, and the penetration of solution in the slices takes time. Therefore, the concentration should be in the range of 5 &#181;M to 25 &#181;M. 2) A 20 &#181;M concentration was selected based on Fried et al.25. Higher concentrations were not tried due to the non-specific toxicity of oligomycin. 3) In the report by Underwood et al.26, the authors did a titration experiment for oligomycin and found that doses at 6.25, 12.5, 25, and 50 &#181;g/mL resulted in similar inhibition. The higher concentration of oligomycin (50 &#181;g/mL) did not inhibit more but had a bigger variance. 4) In our observation, the determining factor seems to be the penetrating ability of oligomycin. It is difficult for oligomycin to penetrate the tissue, and that is why it takes at least 7 to 8 cycles to reach the plateau, the maximum response. As long as it reaches the plateau, the inhibition is assumed to be maximal.
A key technical challenge of adapting the extracellular flux analyzer for measuring OCR in striatal slices is to prevent tissue hypoxia. Since the buffer was not oxygenated during the entire duration of measurements (about 4 h), hypoxia was a central issue. This is especially true for thicker tissue samples, where oxygen cannot diffuse throughout the samples. To overcome this problem, slices were sectioned at 150 &#181;m thickness, so that ambient oxygen could penetrate the middle of the brain slices. In addition, 4 mg/mL bovine serum albumin (BSA) was added to the pre-oxygenated artificial cerebrospinal fluid (ACSF) buffer, which facilitated the determination of maximal OCR, as previously suggested23. We examined whether cells were alive. First, Hoechst 33258 (10 &#181;M) and propidium iodide (10 &#181;M) were used to examine whether cells were healthy under these conditions. We then examined whether medium spiny neurons were functionally healthy using patch-clamp recording. We further evaluated whether dopamine (DA) terminals in the striatal slices were functionally healthy by measuring DA release using fast-scan voltammetry. The results showed that striatal slices that were not oxygenated (ACSF/BSA group) were as healthy as the oxygenated control group24.
We then tested different combinations of slice thickness and punch size to determine optimal striatal slice conditions for the flux respiration assay. Dorsal striatal slices with different thicknesses (150 &#181;m and 200 &#181;m) and punch sizes (1.0 mm, 1.5 mm, and 2.0 mm in diameter) were used for OCR analysis using the analyzer. Striatal slices that were 150 &#181;m thick with a punch size of 1.5 mm in diameter had the highest coupling efficiency and OCRs within an optimal range for the analyzer24.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Accumet AB150 pH benchtop meter</td>
			<td>Thermo Fisher Scientific</td>
			<td>13-636-AB150</td>
			<td>To measure pH</td>
		</tr>
		<tr>
			<td>Antimycin A from streptomyces sp.</td>
			<td>SIGMA</td>
			<td>A8674</td>
			<td>To inhibit complex III of the mitochondria</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>SIGMA</td>
			<td>A6003</td>
			<td>To make modified artificial cerebrospinal fluid (BSA-ACSF)</td>
		</tr>
		<tr>
			<td>Carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP)</td>
			<td>SIGMA</td>
			<td>C2920</td>
			<td>To uncouple mitochondrial respiration</td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>SIGMA</td>
			<td>G8270</td>
			<td>To make artificial cerebrospinal fluid (ACSF)</td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>SIGMA</td>
			<td>D8418</td>
			<td>To dissovle compounds</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>SIGMA</td>
			<td>H3375</td>
			<td>To make artificial cerebrospinal fluid (ACSF)</td>
		</tr>
		<tr>
			<td>Humidified non-CO2 incubator</td>
			<td>Fisher Scientific</td>
			<td>11-683-230D</td>
			<td>To hydrate plates at 37 &#176;C</td>
		</tr>
		<tr>
			<td>Oligomycin from Streptomyces diastatochromogenes</td>
			<td>SIGMA</td>
			<td>O4876</td>
			<td>To inhibit mitochondrial ATP synthase</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>SIGMA-ALDRICH</td>
			<td></td>
			<td>sealing film</td>
		</tr>
		<tr>
			<td>Rotenone</td>
			<td>Tocris</td>
			<td>3616</td>
			<td>To inhibit complex I of the mitochondria</td>
		</tr>
		<tr>
			<td>Seahorse XF Calibrant Solution 500 mL</td>
			<td>Seahorse Bioscience</td>
			<td>103681-100</td>
			<td>Solution for seahorse calibration</td>
		</tr>
		<tr>
			<td>Seahorse XF Extracellular Flux Analyzer</td>
			<td>Seahorse Bioscience</td>
			<td></td>
			<td>Equipment used to analyze oxygen consumption rate, old generation</td>
		</tr>
		<tr>
			<td>Seahorse XFe24 Extracellular Flux Analyzer</td>
			<td>Seahorse Bioscience</td>
			<td></td>
			<td>Equipment used to analyze oxygen consumption rate, new generation</td>
		</tr>
		<tr>
			<td>Seahorse XF24 FluxPaks</td>
			<td>Seahorse Bioscience</td>
			<td>101174-100</td>
			<td>Package of flux analyzer sensor cartridges, tissue culture plates, capture screens,&#160; calibrant solution and calibration plates; assay kit.</td>
		</tr>
		<tr>
			<td>Sodium pyruvate</td>
			<td>SIGMA</td>
			<td>P2256</td>
			<td>To prevent any substrate-limiting constraints of substrate supply</td>
		</tr>
		<tr>
			<td>Stainless steel biopsy punches</td>
			<td>Miltex</td>
			<td></td>
			<td>Device used to punch slices</td>
		</tr>
		<tr>
			<td>Sterile cell culture dish, 35 x 10 mm</td>
			<td>Eppendrof</td>
			<td>0030700102</td>
			<td>Used for slice punch</td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica</td>
			<td>VT1200</td>
			<td>To slice brain tissue</td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>Thermo Scientific Precision</td>
			<td>282-115</td>
			<td>To heat buffer and solutions</td>
		</tr>
	</tbody>
</table>

measurement of oxygen consumption rate in acute striatal slices from adult mice

Mitochondria play an important role in cellular ATP production, reactive oxygen species regulation, and Ca2+ concentration control. Mitochondrial dysfunction has been implicated in the pathogenesis of multiple neurodegenerative diseases, including Parkinson&#39;s disease (PD), Huntington&#39;s disease, and Alzheimer&#39;s disease. To study the role of mitochondria in models of these diseases, we can measure mitochondrial respiration via oxygen consumption rate (OCR) as a proxy for mitochondrial function. OCR has already been successfully measured in cell cultures, as well as isolated mitochondria. However, these techniques are less physiologically relevant than measuring OCR in acute brain slices. To overcome this limitation, the authors developed a new method using a Seahorse XF analyzer to directly measure the OCR in acute striatal slices from adult mice. The technique is optimized with a focus on the striatum, a brain area involved in PD and Huntington&#39;s disease. The analyzer performs a live cell assay using a 24-well plate, which allows the simultaneous kinetic measurement of 24 samples. The method uses circular-punched pieces of striatal brain slices as samples. We demonstrate the effectiveness of this technique by identifying a lower basal OCR in striatal slices of a mouse model of PD. This method will be of broad interest to researchers working in the field of PD and Huntington&#39;s disease.

The first step of this study was to optimize the slice thickness and punch size used to excise a section of the striatum from the slice (Figure 3A). A slice at 150 &#181;m thickness and a 1.5 mm punch size gave the best results determined by the coupling efficiency (Figure 3B-C). As shown in Figure 3B, OCR is relatively stable for 5 h with less than 10% run down. In addition, functional measurements...

Watch this Scientific Journal Video about Measurement of Oxygen Consumption Rate in Acute Striatal Slices from Adult Mice at JoVE.com

Measurement of Oxygen Consumption Rate in Acute Striatal Slices from Adult Mice

Oxygen consumption rate (OCR) is a common proxy for mitochondrial function and can be used to study different disease models. We developed a new method using a Seahorse XF analyzer to directly measure the OCR in acute striatal slices from adult mice that is more physiologically relevant than other methods.

Mitochondria play an important role in cellular ATP production, reactive oxygen species regulation, and Ca2+ concentration control. Mitochondrial ...

measurement-oxygen-consumption-rate-acute-striatal-slices-from-adult

Research

JoVE Journal

Neuroscience

3.1K Views.  Thomas Jefferson University. Oxygen consumption rate (OCR) is a common proxy for mitochondrial function and can be used to study different disease models. We developed a new method using a Seahorse XF analyzer to directly measure the OCR in acute striatal slices from adult mice that is more physiologically relevant than other methods.

Video: Measurement of Oxygen Consumption Rate in Acute Striatal Slices from Adult Mice

Preparation of Acute Hippocampal Slices from Rats and Transgenic Mice for the Study of Synaptic Alterations during Aging and Amyloid Pathology

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The hippocampus can be extracted quickly and easily from rats and mice and slices remain viable for hours in oxygenated artificial cerebrospinal fluid. Moreover, basic electrophysisologic techniques are easily applied to the investigation of synaptic function in hippocampal slices and have provided some of the best biomarkers for cognitive impairments. The hippocampal slice is especially popular for the study of synaptic plasticity mechanisms involved in learning and memory. Changes in the induction of long-term potentiation and depression (LTP and LTD) of synaptic efficacy in hippocampal slices (or lack thereof) are frequently used to describe the neurologic phenotype of cognitively-impaired animals and/or to evaluate the mechanism of action of nootropic compounds. This article outlines the procedures we use for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and Alzheimer's disease (AD)1-3. Use of aged rats and AD model mice can present a unique set of challenges to researchers accustomed to using younger rats and/or mice in their research. Aged rats have thicker skulls and tougher connective tissue than younger rats and mice, which can delay brain extraction and/or dissection and consequently negate or exaggerate real age-differences in synaptic function and plasticity. Aging and amyloid pathology may also exacerbate hippocampal damage sustained during the dissection procedure, again complicating any inferences drawn from physiologic assessment. Here, we discuss the steps taken during the dissection procedure to minimize these problems. Examples of synaptic responses acquired in "healthy" and "unhealthy" slices from rats and mice are provided, as well as representative synaptic plasticity experiments. The possible impact of other methodological factors on synaptic function in these animal models (e.g. recording solution components, stimulation parameters) are also discussed. While the focus of this article is on the use of aged rats and transgenic mice, novices to slice physiology should find enough detail here to get started on their own studies, using a variety of rodent models.

This article outlines procedures for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and age-related neurodegenerative diseases, such as Alzheimer&#x2019;s disease.

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The ...

Presynaptic Dopamine Dynamics in Striatal Brain Slices with Fast-scan Cyclic Voltammetry

Extensive research has focused on the neurotransmitter dopamine because of its importance in the mechanism of action of drugs of abuse (e.g. cocaine and amphetamine), the role it plays in psychiatric illnesses (e.g. schizophrenia and Attention Deficit Hyperactivity Disorder), and its involvement in degenerative disorders like Parkinson's and Huntington's disease. Under normal physiological conditions, dopamine is known to regulate locomotor activity, cognition, learning, emotional affect, and neuroendocrine hormone secretion. One of the largest densities of dopamine neurons is within the striatum, which can be divided in two distinct neuroanatomical regions known as the nucleus accumbens and the caudate-putamen. The objective is to illustrate a general protocol for slice fast-scan cyclic voltammetry (FSCV) within the mouse striatum. FSCV is a well-defined electrochemical technique providing the opportunity to measure dopamine release and uptake in real time in discrete brain regions. Carbon fiber microelectrodes (diameter of ~ 7 &mu;m) are used in FSCV to detect dopamine oxidation. The analytical advantage of using FSCV to detect dopamine is its enhanced temporal resolution of 100 milliseconds and spatial resolution of less than ten microns, providing complementary information to in vivo microdialysis.

Using fast-scan cyclic voltammetry to measure electrically evoked presynaptic dopamine dynamics in striatal brain slices.

Extensive research has focused on the neurotransmitter dopamine because of its importance in the mechanism of action of drugs of abuse (e.g. cocaine and ...

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 ...

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with enhanced [Ca2+]i signals. It has been proposed that astrocytes in turn regulate neurons and blood vessels through calcium-dependent mechanisms, such as the release of signaling molecules. However, [Ca2+]i imaging in entire astrocytes has only recently become feasible with genetically encoded calcium indicators (GECIs) such as the GCaMP series. The use of GECIs in astrocytes now provides opportunities to study astrocyte [Ca2+]i signals in detail within model microcircuits such as the striatum, which is the largest nucleus of the basal ganglia. In the present report, detailed surgical methods to express GECIs in astrocytes in vivo, and confocal imaging approaches to record [Ca2+]i signals in striatal astrocytes in situ, are described. We highlight precautions, necessary controls and tests to determine if GECI expression is selective for astrocytes and to evaluate signs of overt astrocyte reactivity. We also describe brain slice and imaging conditions in detail that permit reliable [Ca2+]i imaging in striatal astrocytes in situ. The use of these approaches revealed the entire territories of single striatal astrocytes and spontaneous [Ca2+]i signals within their somata, branches and branchlets. The further use and expansion of these approaches in the striatum will allow for the detailed study of astrocyte [Ca2+]i signals in the striatal microcircuitry.

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ.

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with ...

Investigation of Synaptic Tagging/Capture and Cross-capture using Acute Hippocampal Slices from Rodents

Synaptic tagging and capture (STC) and cross-tagging are two important mechanisms at cellular level that explain how synapse-specificity and associativity is achieved in neurons within a specific time frame. These long-term plasticity-related processes are the leading candidate models to study the basis of memory formation and persistence at the cellular level. Both STC and cross-tagging involve two serial processes: (1) setting of the synaptic tag as triggered by a specific pattern of stimulation, and (2) synaptic capture, whereby the synaptic tag interacts with newly synthesized plasticity-related proteins (PRPs). Much of the understanding about the concepts of STC and cross-tagging arises from the studies done in CA1 region of the hippocampus and because of the technical complexity many of the laboratories are still unable to study these processes. Experimental conditions for the preparation of hippocampal slices and the recording of stable late-LTP/LTD are extremely important to study synaptic tagging/cross-tagging. This video article describes the experimental procedures to study long-term plasticity processes such as STC and cross-tagging in the CA1 pyramidal neurons using stable, long-term field-potential recordings from acute hippocampal slices of rats.

This video article describes experimental procedures to study long-term plasticity and its associative processes such as synaptic tagging, capture and cross-tagging in the CA1 pyramidal neurons using acute hippocampal slices from rodents.

Synaptic tagging and capture (STC) and cross-tagging are two important mechanisms at cellular level that explain how synapse-specificity and associativity ...

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding diseases, and future therapies hinge on fundamental understandings about how different retinal cells function normally. Gaining such information with biochemical methods has proven difficult because contributions of particular cell types are diminished in the retinal cell milieu. Live retinal imaging can provide a view of numerous biological processes on a subcellular level, thanks to a growing number of genetically encoded fluorescent biosensors. However, this technique has thus far been limited to tadpoles and zebrafish larvae, the outermost retinal layers of isolated retinas, or lower resolution imaging of retinas in live animals. Here we present a method for generating live ex vivo retinal slices from adult zebrafish for live imaging via confocal microscopy. This preparation yields transverse slices with all retinal layers and most cell types visible for performing confocal imaging experiments using perfusion. Transgenic zebrafish expressing fluorescent proteins or biosensors in specific retinal cell types or organelles are used to extract single-cell information from an intact retina. Additionally, retinal slices can be loaded with fluorescent indicator dyes, adding to the method&#39;s versatility. This protocol was developed for imaging Ca2+ within zebrafish cone photoreceptors, but with proper markers it could be adapted to measure Ca2+ or metabolites in M&#252;ller cells, bipolar and horizontal cells, microglia, amacrine cells, or retinal ganglion cells. The retinal pigment epithelium is removed from slices so this method is not suitable for studying that cell type. With practice, it is possible to generate serial slices from one animal for multiple experiments. This adaptable technique provides a powerful tool for answering many questions about retinal cell biology, Ca2+, and energy homeostasis.

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

Imaging retinal tissue can provide single-cell information that cannot be gathered from traditional biochemical methods. This protocol describes preparation of retinal slices from zebrafish for confocal imaging. Fluorescent genetically encoded sensors or indicator dyes allow visualization of numerous biological processes in distinct retinal cell types.

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding ...

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.