Funding provided by the National Institute on Alcohol Abuse and Alcoholism (NIAAA; AA-016967 and AA016967-01S1; TAM), Wayne State University start up funds, and Wayne State University Research Grant Program. The content is solely the responsibility of the authors and does not represent the official views of NIAAA or the National Institutes of Health.

1. Experimental essentials
Electrode Fabrication
<ul type="circle">
 <li>There are numerous carbon fiber microelectrodes fabrication methods since most are made in-house. Typically what dictates the electrode fabrication details is the electrochemical technique that is applied to the electrode (e.g. amperometry vs. FSCV). For FSCV, microelectrodes can be made in-house using the following three-step procedure. For a more complete description of carbon fiber electrode fabrication, see a recent JOVE article 1. However, note that the electrodes described below are cylindrical carbon fiber microelectrodes, which require fewer steps to fabricate versus the amperometric carbon fiber microelectrodes from the above-mentioned protocol. This simplified protocol does not require boiling the carbon fiber in acetone, fire-polishing the glass capillaries, or using epoxy to seal the glass-fiber junction.</li>
 <li>Using vacuum suction aspirate a carbon fiber (diameter 7 &mu;m; Goodfellow Oakdale, PA) into a borosilicate glass capillary with microfilament (length 10 cm, o.d. 1.2 mm, i.d. 0.68 mm; A-M systems, Carlsborg, WA).</li>
 <li>Place the threaded capillary into the electrode puller (Narishige, Tokyo, Japan) where the capillary is pulled in half. Output settings for the electrode puller do vary from lab to lab. For reference, our output settings for the puller are 90.7 main magnet, 23.2 sub-magnet, and 53.4 for the heater. The output settings should be empirically refined to generate a glass taper that is approximately 4.4 mm in length, with a tight seal around the carbon fiber.</li>
 <li>Under a microscope (Olympus, Tokyo, Japan), trim the carbon fiber (using a scalpel blade) extending from the glass tip allowing approximately 50-200 &mu;m of the carbon fiber to protrude from the tightly sealed interface.</li>
</ul>
Solution Preparation
Three types of artificial cerebrospinal fluid (aCSF) solutions need to be prepared in advance, all in ultrapure (18 M&Omega; cm) water.
<ul type="circle">
 <li>Sucrose-aCSF can be prepared at least one day before the slicing. The sucrose-aCSF buffer consists of (in mM): 180 sucrose, 30 NaCl, 4.5 KCl, 1.0 MgCl2, 26 NaHCO3, 1.2 NaH2PO4, and 10 D-glucose (pH 7.4) and should be oxygenated using 95% O2/5% CO2 for 15 minutes 2. If prepared ahead of time, the solution can be kept refrigerated at 4 &deg;C for up to 1 week.</li>
 <li>ACSF solution for voltammetric recordings should be prepared the day of the experiment. The aCSF solution consists of (in mM): 126 NaCl, 2.5 KCl, 2.4 CaCl2, 1.2 MgCl2, 25 NaHCO3, 1.2 NaH2PO4, 11 D-glucose, 0.4 ascorbic acid (pH 7.4). During the course of the experiment, oxygenate by bubbling with 95% O2/5% CO2 at room temperature.</li>
 <li>Modified-aCSF solution for electrode calibrations contains (in mM): 2.5 KCl, 126 NaCl, 1.2 NaH2PO4, 2.4 CaCl2, 1.2 MgCl2, and 25 NaHCO3 (pH 7.4) and can be kept for up to one week, without refrigeration or oxygenation.</li>
</ul>
Electrode Calibration
<ul type="circle">
 <li>Electrode viability and sensitivity is determined by pre- and post-calibration, respectively, using a flow "t-cell" containing 3 ports with a sealed reference electrode (Ag/AgCl). The fabricated microelectrode is lowered into the flow "t-cell". The inlet port is connected to a syringe pump, allowing for a continuous flow of modified-aCSF at a flow rate of 2 mL/min. The third port of the "t-cell" is connected to a syringe filled with 3 &mu;M dopamine, made in modified-aCSF.</li>
 <li>To pre-calibrate, allow modified aCSF solution to flow for about 3 - 7 seconds, and then manually inject 1-2 mL of the 3 &mu;M dopamine standard. For each electrode to be pre-calibrated, repeat the calibration at least 3 times and average the maximal currents obtained from each.</li>
 <li>Immediately after the slice voltammetry experiment (see section 3), the electrode is post-calibrated in the same manner as described above.</li>
 <li>The calibration factor (later used during data analysis) is determined by dividing the average dopamine oxidation current (nA) by the concentration of the dopamine standard. For example, the current response is divided by 3 when using a 3 &mu;M dopamine standard.</li>
</ul>
2. Slice preparation
<ol>
 <li>Pour 10 mL of sucrose aCSF into a small beaker and place in an ice bucket. Additionally, place instant adhesive (Loctite 404) in ice bucket, within reach.</li>
 <li>Prepare a razor blade and the necessary tools required for dissection, such as forceps, spatula, and scissors, by wiping them clean with an alcohol pad.</li>
 <li>Sacrifice mouse by CO2 asphyxiation in a small gas chamber, followed by immediate decapitation using sharp scissors. Quickly remove the entire brain. Place the brain in beaker of ice-cold sucrose aCSF for approximately 10 minutes.</li>
 <li>In the meantime, prepare Vibratome for slicing. First, place some crushed ice in the specimen bath. Position the specimen chamber and tighten to hold it firmly in place. Add more ice around the specimen chamber to fill in the gaps, making sure no ice gets into the chamber. Place the cleaned razor blade in the blade holder on the Vibratome and fill the specimen chamber with ice-cold sucrose aCSF.</li>
 <li>To set up a working surface for preparing the brain, pour some cold sucrose aCSF on a piece of paper towel that has been placed on an upturned Petri dish. Using forceps transfer the brain onto the prepared Petri dish. For coronal slices, cut the cerebellum along the medial-lateral axis with a razor blade and discard. This creates a flat base that can be affixed to the Vibratome stage.</li>
 <li>Place a drop of the Loctite adhesive (placed in the ice bucket during Step 1) on the specimen stage. Immediately, affix the flat end of the prepared brain on the stage, keeping it as upright as possible. Place the stage in the specimen chamber and tighten the screw, ensuring that the brain is completely immersed in the sucrose aCSF in the chamber.</li>
 <li>Using the controls on the front of the Vibratome adjust the stage so that the razor blade lines up with the top of the brain. Optimal parameters for the Vibratome are obtained by setting the frequency and speed to low. Lower speeds are preferred to minimize the force of the blade from squashing the brain, which is observed at higher speeds. Slice thickness is set to 400 &mu;m.</li>
 <li>The first few slices will not contain the striatum. Repeat the slicing until slices containing the striatum are obtained. Once the striatal region is reached (affirmed by anatomical landmarks), use a paint brush to lift the slice and place in a beaker with oxygenated, room-temperature aCSF. Typically, one can obtain 3 to 4 slices containing the striatal complex so that the caudate-putamen and nucleus accumbens are included.</li>
 <li>Allow the slices to acclimate in oxygenated aCSF at room temperature for at least 1 hour before using for experiments.</li>
</ol>
3. Voltammetric recordings from slices
While the slices are incubating, the slice recording chamber can be prepared.
<ol>
 <li>Take tubing connected to the submersion recording chamber (Custom Scientific, Denver, CO) and place in oxygenating, room temperature aCSF. Set the perfusion pump (Watson Marlow Limited, Falmouth, England) to a flow rate of 1 mL/min. Set the temperature controller to 32 &deg;C. After aCSF fills the custom-built slice holder (modified mesh disc stage), prepare it for the slice by removing any air bubbles using a needle syringe (BD spinal needle).</li>
 <li>Prime the slice bath by drawing a vacuum on the outflow tubing (leading to glass liquid waste bottle) using a syringe to start flow. Place a kimwipe to act as a wick on the edge of the slice holder to control buffer overflow.</li>
 <li>After 1 hour incubation, transfer the slices to the slice holder in the recording chamber, which is continuously perfused with 95% O2/5% CO2 room temperature aCSF.</li>
 <li>Submerge the Ag/AgCl reference electrode in the slice holder (taped to lid of slice chamber) and connect the electrode using an alligator clip to the head stage.
 <ul type="circle">
 <li>The Ag/AgCl reference electrode can be made in house by anodizing (+1 V) a 250 &mu;m silver wire (A-M Systems, Carlsborg, WA) in 1 M HCl for 5 minutes to deposit a thin layer of AgCl on the surface of the silver wire.</li>
 </ul>
 </li>
 <li>Lower the tungsten stimulating electrode (Plastics One, Roanoke, VA) to the surface of the striatal brain slice. The stimulating electrode makes contact with the slice as it rests on top of it, but should not puncture the slice. In our experimental set-up, the stimulation is generated by a Neurolog stimulator.</li>
 <li>The carbon fiber working microelectrode is back-filled with a 150 mM KCl solution using a BD spinal needle. A lead wire is then inserted (Squires Electronics, Cornelius, OR), which is connected to the head stage of the low-noise ChemClamp potentiostat (Dagan Corporations, Minneapolis, MN) using an alligator clip. The working electrode is placed approximately 100 - 200 &mu;m away from the bipolar stimulating electrodes, about 75 &mu;m deep into the slice.</li>
 <li>Electrical stimulations, using either single (monophasic, 350 &mu;A, 60 Hz, and 4 ms pulse width) or multiple (e.g. 5 pulses, 350 &mu;A, 20 Hz, and 4 ms wide) pulses, are delivered by the stimulating electrode to evoke neurotransmitter release 3.</li>
 <li>To measure electrically evoked dopamine using FSCV, a triangle waveform is applied to the electrode. Typical parameters for dopamine detection: potential of the carbon fiber microelectrode is held at -0.4 V versus a Ag/AgCl reference electrode, ramped to a positive limit of +1.2 V, then brought back down to -0.4 V at a scan rate of 400 V/s.</li>
 <li>The slice is electrically stimulated every 5 minutes and voltammetric measurements of the resulting dopamine efflux are made for 15 seconds.</li>
 <li>After at least three stable electrically stimulated dopamine release recordings (difference between peak height is &lt; 10%), aCSF containing pharmacological agent of interest is perfused at a flow rate of 1 mL/min over the slice for 30 minutes to obtain maximal effect. Electrically stimulated dopamine recordings are made every 5 minutes during the pharmacological perfusion.</li>
</ol>
4. Data analysis
Resulting current versus time traces obtained from the slice can be fit by nonlinear regression to a set of Michaelis-Menten based equations, as described by Wightman and colleagues in software written in LabVIEW (National Instruments, Austin, TX) 4-6. In this software, current versus time traces can be fitted by varying two parameters, Vmax (nM/s; corresponding to the rate of uptake by the dopamine transporter as demonstrated by the descending phase of the trace) and dopamine concentration per pulse (nM; corresponding to the peak height maximum). The Km value, reflective of the affinity of dopamine for the dopamine transporter, is set to 160 nM and not changed. The electrode's post-calibration factor that is determined after the experiment is required prior to fitting. The LabView software contains the coefficient of determination (R2) parameter to determine the goodness of fit (R2 values &gt; 0.8 are used).
5. Representative Results
FSCV was used to examine single-pulse, electrically stimulated dopamine release and uptake in the caudate-putamen (CPu), nucleus accumbens (NAc) core, and NAc shell in mice. Representative results shown in Figure 1A demonstrate current (or concentration) versus time plots. The red arrow indicates when electrical stimulation is applied to the slice followed by a corresponding rise in the amount of current attributable to changes in dopamine concentration at the carbon fiber microelectrode. The predominant process during electrical stimulation is dopamine release, but other processes such as uptake and diffusion contribute to the overall observed peak height as well. The descending phase of the peak is mainly attributed to reuptake of the neurotransmitter by its transporter since neuronal stimulation has been stopped 7. However, the peak decay is not limited to reuptake, as diffusion and metabolism also contribute to the decrease in current. It has been postulated that since the time scale of electrochemical measurements is a matter of seconds, FSCV is too fast to measure contributions from metabolism 7. In these current versus time traces, the y-axis is converted to concentration (&mu;M) using the post-experiment calibration factor. The inset for Figure 1A is the respective background subtracted cyclic voltammograms for the current versus time traces. Plotting the measured current (y-axis) against the applied potential to the electrode (x-axis), dopamine is chemically identified with an observed oxidation peak at +0.6 V and a corresponding reduction peak of dopamine-ortho-quinone is observed at -0.2 V versus a Ag/AgCl reference electrode. The third representation of the data uses a three-dimensional pseudo-color plot (Figure 1B) combining the information from both the current versus time traces and the cyclic voltammogram to form a single plot. In the representative pseudo-color plot, time is plotted in seconds along the x-axis, the applied voltage to the carbon fiber working microelectrode is graphed along the y-axis, and current is represented as false color along the z-axis. Due to the small size of carbon fiber microelectrodes (~ 7 &mu;m in diameter), electrically stimulated dopamine dynamics can be detected in discrete anatomical regions of the striatum (CPu versus NAc core versus NAc shell; Figure 2).
An advantage of using striatal coronal slices is that it eliminates contributions from the dopamine cell bodies, and allows for an investigation of presynaptic dopamine dynamics. Presynaptic control of dopamine release and uptake is not strictly limited to dopamine autoreceptor or transporter functions as others have shown 16, 17. Heteroreceptors of other neurotransmitter systems also modulate dopamine dynamics 18, 19. Representative current versus time traces shown in Figure 3A demonstrate that when slices are treated with 1 &mu;M quinpirole (D2/D3 receptor agonist) for 30 minutes, a decrease in electrically-evoked dopamine release is observed. On the other hand, when a substrate for the dopamine transporter, such as methamphetamine, is perfused over the slice for 30 minutes, no difference in dopamine release is observed (Figure 3B). The peak decay is shifted to the right, which is typically associated with alterations in dopamine transporter kinetics (Km) 3. Finally, Figure 3C is a representative trace of the current versus time trace once the slice has been bathed in a 100 ng/mL brain-derived neurotrophic factor (BDNF) solution, which has been hypothesized to influence dopamine release dynamics 20, 21. From this representative trace, it can be seen that BDNF has the ability to enhance electrically-evoked dopamine release. Taken together, these pharmacological treatments emphasize the utility of FSCV to probe dopamine dynamics within the striatum.
The primary limitation of using brain slices to investigate presynaptic dopamine dynamics by FSCV is that the neurocircuitry from an intact brain is lost. With slice FSCV it is impossible to study the effects of neurotransmitters from other brain regions, making it difficult to understand contributions of these systems on the functionality of the region being investigated (e.g. the striatum) or to evaluate non-stimulated dopamine levels. However, recent technical advances in FSCV has allowed for dopamine transient measurements (with and without electrical stimulation) in freely moving rats in response to a pharmacological manipulation, self-administration, or novelty 22 - 24. Overall, slice FSCV provides valuable information on presynaptic dopamine dynamics, and coupling slice FSCV results to complementary neurochemical techniques such as microdialysis, electrophysiology, and/or freely moving FSCV offers a more comprehensive view of neurotransmitter functioning in the brain.
<img alt="Figure 1" src="/files/ftp_upload/3464/3464fig1.jpg" /> 
Figure 1. Electrically-evoked dopamine release measured using FSCV following single-pulse stimulation in dorsal CPu slices from C57Bl/6J mice. (A) The concentration versus time trace in which dopamine release was evoked by a single pulse (red arrow). The single asterisk represents the factors that contribute to the rise in concentration, which is predominantly dopamine release, but uptake and diffusion also contribute. The double asterisk represents the peak signal returning to baseline, mainly due to uptake but also diffusion contributes. Inset displays the corresponding cyclic voltammograms. (B) Representative color plots from the dorsal CPu display time (x-axis), applied potential to the carbon fiber microelectrode versus Ag/AgCl reference electrode (y-axis), and current in pseudo-color.
<img alt="Figure 2" src="/files/ftp_upload/3464/3464fig2.jpg" /> 
Figure 2. Dopamine release evoked by a single electrical stimulation pulse (indicated by the red arrow) in the NAc core and shell from C57Bl/6J mice. (A and C) The concentration versus time traces and their corresponding cyclic voltammograms (inset) from the NAc core and shell. (B and D) As previously described, representative color plots from the NAc core and shell.
<img alt="Figure 3" src="/files/ftp_upload/3464/3464fig3.jpg" /> 
Figure 3. Representative traces after the slice was treated with a pharmacological agent for 30 minutes; in all cases, a single pulse was used to evoke dopamine release in the CPu. (A) Application of the dopamine D2 receptor agonist, quinpirole (red trace) compared to pre-treatment (black trace). (B) Methamphetamine perfusion (purple trace) compared to pre-treatment (black trace). (C) The ability of brain-derived neurotrophic factor to influence dopamine dynamics (blue trace) compared to pre-treatment (black trace).

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No conflicts of interest declared.

The protocol presented here demonstrates how to prepare and use mouse coronal brain slices for FSCV experiments. Although this method is specific to obtaining and measuring dopamine dynamics, other neurotransmitters such as adenosine, hydrogen peroxide, norepinephrine, and serotonin have been monitored in vivo or in vitro with FSCV 3, 8 - 11. FSCV can be used to monitor some of these other neurochemicals by simple modifications of the waveform applied to the working electrode 3, 11...

<table border="1">
<tbody>
<tr>
<td>Name</td>
<td>Company</td>
<td>CAS number</td>
</tr>
<tr>
<td width="274">Reagent</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Potassium Chloride</td>
<td>Fisher </td>
<td width="142" valign="top">7447407</td>
</tr>
<tr>
<td width="274">Sodium chloride</td>
<td>EMD Chemicals</td>
<td width="142" valign="top">7647145</td>
</tr>
<tr>
<td width="274">Magnesium chloride</td>
<td width="194" valign="top">Fisher</td>
<td width="142" valign="top">7791186</td>
</tr>
<tr>
<td width="274">Calcium chloride</td>
<td width="194" valign="top">Fisher</td>
<td width="142" valign="top">10035048</td>
</tr>
<tr>
<td width="274">Sodium bicarbonate </td>
<td width="194" valign="top">EMD Chemicals</td>
<td width="142" valign="top">144558</td>
</tr>
<tr>
<td width="274">Sodium phosphate,Dibasic</td>
<td width="194" valign="top">EMD Chemicals</td>
<td width="142" valign="top">7558794</td>
</tr>
<tr>
<td width="274">D-glucose</td>
<td width="194" valign="top">Fisher</td>
<td width="142" valign="top">50997</td>
</tr>
<tr>
<td width="274">Ascorbic acid</td>
<td width="194" valign="top">Fisher</td>
<td width="142" valign="top">50817</td>
</tr>
<tr>
<td width="274">Sucrose</td>
<td width="194" valign="top">Fisher</td>
<td>57501</td>
</tr>
<tr>
<td width="274">&nbsp;</td>
<td width="194" valign="top">&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Materials</td>
<td width="194" valign="top">Vendor</td>
<td>Catalogue number </td>
</tr>
<tr>
<td width="274">Carbon fiber</td>
<td>Goodfellow Oakdale, PA</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Glass capillary </td>
<td>A-M Systems Carlsborg&nbsp; WA.</td>
<td>602000</td>
</tr>
<tr>
<td width="274">Silver wire</td>
<td>A-M Systems Carlsborg&nbsp; WA.</td>
<td>787000</td>
</tr>
<tr>
<td width="274">Tungsten stimulating electrode </td>
<td>Plastics One, Roanoke, VA </td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Platinum wire </td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Lead wire</td>
<td>Squires Electronics, Cornelius, OR</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Loctite 404 instant adhesive </td>
<td>Hankal Corp. Rocky Hill 
CT.</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Razor blade</td>
<td>World precision Instruments Inc. FL.</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">BD Spinal needle</td>
<td>BD Medical systems, 
Franlin&nbsp; Lake, NJ</td>
<td>REF 405234</td>
</tr>
<tr>
<td width="274">Surgical Blade </td>
<td>Feather&nbsp; Safety razor Co. LTD. Japan</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Software </td>
<td width="194" valign="top">Vendor</td>
<td>Catalogue number </td>
</tr>
<tr>
<td width="274">TH software</td>
<td>ESA Inc.,Chelmsford, MA</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Instrument</td>
<td width="194" valign="top">Vendor</td>
<td>Catalogue number </td>
</tr>
<tr>
<td width="274">Submersion recording chamber </td>
<td>Custom Scientific, Denver, CO </td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Neorolog stimulus isolator</td>
<td>Digitmeter, Hertfordshire, England </td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Automatic&nbsp; temperature controller</td>
<td>Warner Instrument Corporation </td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Microscope (SZX7)</td>
<td>Olympus</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Microscope </td>
<td>Fisher</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Vibratome 3000 sectioning&nbsp; system</td>
<td>St. Louis , MO.</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">Perfusion pump </td>
<td>Watson Marlow Limited, Falmouth Cornwall &ndash; England</td>
<td>H110708</td>
</tr>
<tr>
<td width="274">Micropipette puller</td>
<td>Narishige, Tokyo, Japan</td>
<td>&nbsp;</td>
</tr>
<tr>
<td width="274">ChemClamp potentiostat </td>
<td>Dagan Corporations, 
Minneapolis, MN. </td>
<td>&nbsp;</td>
</tr>
</tbody>
</table>

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.

Watch this Scientific Journal Video about Presynaptic Dopamine Dynamics in Striatal Brain Slices with Fast-scan Cyclic Voltammetry at JoVE.com

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

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 ...

presynaptic-dopamine-dynamics-striatal-brain-slices-with-fast-scan

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Neuroscience

21.7K Views.  Wayne State University. Using fast-scan cyclic voltammetry to measure electrically evoked presynaptic dopamine dynamics in striatal brain slices.

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

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

Voltage Biasing, Cyclic Voltammetry, &amp; Electrical Impedance Spectroscopy for Neural Interfaces

Electrical impedance spectroscopy (EIS) and cyclic voltammetry (CV) measure properties of the electrode-tissue interface without additional invasive procedures, and can be used to monitor electrode performance over the long term. EIS measures electrical impedance at multiple frequencies, and increases in impedance indicate increased glial scar formation around the device, while cyclic voltammetry measures the charge carrying capacity of the electrode, and indicates how charge is transferred at different voltage levels. As implanted electrodes age, EIS and CV data change, and electrode sites that previously recorded spiking neurons often exhibit significantly lower efficacy for neural recording. The application of a brief voltage pulse to implanted electrode arrays, known as rejuvenation, can bring back spiking activity on otherwise silent electrode sites for a period of time. Rejuvenation alters EIS and CV, and can be monitored by these complementary methods. Typically, EIS is measured daily as an indication of the tissue response at the electrode site. If spikes are absent in a channel that previously had spikes, then CV is used to determine the charge carrying capacity of the electrode site, and rejuvenation can be applied to improve the interface efficacy. CV and EIS are then repeated to check the changes at the electrode-tissue interface, and neural recordings are collected. The overall goal of rejuvenation is to extend the functional lifetime of implanted arrays.

The electrode-tissue interface of neural recording electrodes can be characterized with electrical impedance spectroscopy (EIS) and cyclic voltammetry (CV). Application of voltage biasing changes the electrochemical properties of the electrode-tissue interface and can improve recording capability. Voltage biasing, EIS, CV, and neural recordings are complementary.

Electrical impedance spectroscopy (EIS) and cyclic voltammetry (CV) measure properties of the electrode-tissue interface without additional invasive ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Modeling Fast-scan Cyclic Voltammetry Data from Electrically Stimulated Dopamine Neurotransmission Data Using QNsim1.0

Central dopaminergic (DAergic) pathways have an important role in a wide range of functions, such as attention, motivation, and movement. Dopamine (DA) is implicated in diseases and disorders including attention deficit hyperactivity disorder, Parkinson's disease, and traumatic brain injury. Thus, DA neurotransmission and the methods to study it are of intense scientific interest. In vivo fast-scan cyclic voltammetry (FSCV) is a method that allows for selectively monitoring DA concentration changes with fine temporal and spatial resolution. This technique is commonly used in conjunction with electrical stimulations of ascending DAergic pathways to control the impulse flow of dopamine neurotransmission. Although the stimulated DA neurotransmission paradigm can produce robust DA responses with clear morphologies, making them amenable for kinetic analysis, there is still much debate on how to interpret the responses in terms of their DA release and clearance components. To address this concern, a quantitative neurobiological (QN) framework of stimulated DA neurotransmission was recently developed to realistically model the dynamics of DA release and reuptake over the course of a stimulated DA response. The foundations of this model are based on experimental data from stimulated DA neurotransmission and on principles of neurotransmission adopted from various lines of research. The QN model implements 12 parameters related to stimulated DA release and reuptake dynamics to model DA responses. This work describes how to simulate DA responses using QNsim1.0 and also details principles that have been implemented to systematically discern alterations in the stimulated dopamine release and reuptake dynamics.

Fast-scan cyclic voltammetry can monitor in vivo dopamine neurotransmission in the context of drugs, disease, and other experimental manipulations. This work describes the implementation of QNsim1.0, a software to model electrically stimulated dopamine responses according to the quantitative neurobiological model to quantify estimates of dopamine release and reuptake dynamics.

Central dopaminergic (DAergic) pathways have an important role in a wide range of functions, such as attention, motivation, and movement. Dopamine (DA) is ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Stereotaxic Surgery for Genetic Manipulation in Striatal Cells of Neonatal Mouse Brains

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, they may function to regulate the developmental process and/or physiological function in neonatal brains. To investigate neurobiological functions of specific genes in the brain, it is essential to inactivate genes in the brain. Here, we describe a simple stereotaxic method to inactivate gene expression in the striatum of transgenic mice at neonatal time windows. AAV-eGFP-Cre viruses were microinjected into the striatum of Ai14 reporter gene mice at postnatal day (P) 2 by stereotaxic brain surgery. The tdTomato reporter gene expression was detected in P14 striatum, suggesting a successful Cre-loxP mediated DNA recombination in AAV-transduced striatal cells. We further validated this technique by microinjecting AAV-eGFP-Cre viruses into P2Foxp2fl/fl mice. Double labeling of GFP and Foxp2 showed that GFP-positive cells lacked Foxp2 immunoreactivity in P9 striatum, suggesting the loss of Foxp2 protein in AAV-eGFP-Cre transduced striatal cells. Taken together, these results demonstrate an effective genetic deletion by stereotaxically microinjected AAV-eGFP-Cre viruses in specific neuronal populations in the neonatal brains of floxed transgenic mice. In conclusion, our stereotaxic technique provides an easy and simple platform for genetic manipulation in neonatal mouse brains. The technique can not only be used to delete genes in specific regions of neonatal brains, but it also can be used to inject pharmacological drugs, neuronal tracers, genetically modified optogenetics and chemogenetics proteins, neuronal activity indicators and other reagents into the striatum of neonatal mouse brains.

We describe a protocol of stereotaxic surgery with a homemade head-fixed device for microinjecting reagents into the striatum of neonatal mouse brains. This technique allows genetic manipulation in neuronal cells of specific regions of neonatal mouse brains.

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, ...

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 ...