This work was supported by grants of the Deutsche Forschungsgemeinschaft through the Collaborative Research Center 889 &#34;Cellular Mechanisms of Sensory Processing&#34; to S.T. (Project C04). We thank Sina Pl&#252;mer, Leonore Burchardt, Dirk Pr&#252;sse, Klaus Heisig and Ralf Brockhausen for technical and animal-related support and our collaborators in the Stem Cell Unit of the German Primate Center, Dr. Katharina Debowski and Anna Magerhans, for technical assistance in the filtration process.

Animal care and all experimental procedures were conducted in accordance with German laws governing animal care and approved by the district government of Braunschweig, Lower Saxony, Germany.
Note: As the experiment is performed in vivo, it is crucial to maintain the highest possible hygiene standards. Whenever possible, work under sterile conditions.
1. Preparing the Injection/Recording System
<ol>
	<li>Sterilize the tube that connects the micropipette with the syringe. Use the shortest length of tube possible in the experimental set-up between injection pump and electrophysiological recording system.</li>
	<li>Clean the guide tubes of the recording system using cleaning wires. Dip them in sterile silicon oil and feed them through the individual guide tubes several times.</li>
	<li>Insert the quartz glass micropipette into one guide tube of the recording system. See Figure 1A.</li>
	<li>After fixing the micropipette in the recording system, attach the sterile tube to the metal pin of the micropipette. Take care; although the micropipette is fixed in the system it can easily break when attaching the tube. Use two sterile tweezers to apply equal pressure on pin and tube.</li>
	<li>Use liquid super glue to seal the junction between tube and micropipette. Wait at least 3 hr for the glue to harden before filling the micropipette with liquid.</li>
	<li>Insert microelectrodes (e.g., quartz glass insulated platinum tungsten) into the other positions of the recording system before or after micropipette insertion.</li>
</ol>
2. Preparing the Substance
<ol>
	<li>Sterilize 1.5 ml microcentrifuge tubes for later storage of injection solutions using an autoclave or other reliable procedure.</li>
	<li>Weigh the corresponding substance scopolamine hydrochloride to prepare 5&#160;ml of a 0.1 molar solution. Dissolve in sterile saline (0.9% NaCl).</li>
	<li>Under sterile conditions in a fume hood, filter the solution using a syringe filter with sufficiently large pore diameter, e.g., 0.2 &#181;m.</li>
	<li>Under the fume hood, aliquot the solution into volumes sufficient for a single experiment, e.g., for scopolamine, 500 &#181;l in sterile 1.5 ml microcentrifuge tube. Use dark tubes to protect the substance from light; alternatively, wrap tubes in aluminum foil. For scopolamine, store the solution for up to 14 days at 4 &#176;C.</li>
</ol>
3. Daily Preparation of the Injection/Recording System
Note: When mounted in the recording system, the electrodes and micropipette are stored in an enzyme solution (Tergazyme, 1% solution with deionized water) between recordings. The following steps must be performed before every recording.
<ol>
	<li>Collect a tube of the substance to be injected. Allow it to reach RT if refrigerated.</li>
	<li>Remove the injection/recording system from the enzyme solution and rinse electrodes and micropipette with deionized water to clean completely of enzyme solution.</li>
	<li>Remove the front cover of the recording system in order to visually check the seal between micropipette and tube.</li>
	<li>Apply sterile silicon oil to the guide tube gap (see Figure 1A) and tips of electrodes and micropipette in order to lubricate the system for smooth movement.</li>
	<li>Check tips of electrodes and micropipette using a microscope to ensure they are intact. Align the electrodes and the micropipette under the microscope so that they extend out of the guide tubes with the same length. Drive them into the guide tubes, stopping as soon as they are no longer visible. This is defined as electrode position zero. Set the depth of the electrodes and micropipette to 0 in the software.</li>
	<li>Fill a sterile syringe with sterile saline and insert the needle into the tube, taking care not to pierce the wall of the tube. Drive the micropipette out of the guide tube for visual control of substance flow.</li>
	<li>Flush at least 2 ml sterile saline through the tube and micropipette to ensure no air remains in the syringe or in the tube. Do not apply too much pressure to the plunger of the syringe. Ensure the junction between tube and micropipette is sealed. If leakage is visible, re-glue the junction (see step 1.5) and postpone recording.</li>
	<li>Fill a new sterile syringe with the solution to be injected and exchange it with the barrel of the saline syringe, i.e., keep the needle of the saline-filled syringe in the tube. Make sure that no air is transferred into the system. This is best achieved by filling the needle hub with saline after removing the saline-filled barrel.</li>
	<li>Flush the system with 250 &#181;l of the solution to be injected, in order to completely remove the saline from the tube.</li>
	<li>Using the motor control software, retract electrodes and micropipette into the guidetubesto a depth of at least -500 &#181;m.</li>
	<li>Lower the guide tube ring (Figure 1A) to the bottom of the guide tubes to maintain their fixed relative position.</li>
	<li>Clean the base of the system with ethanol, in particular where it will touch the monkey&#39;s recording chamber.</li>
	<li>Close the recording system by replacing the front cover and tightening the screws.</li>
</ol>
4. Validation of the Injection System
Note: Although the company calibrates the system, it is recommended to validate the ejected volumes with the materials used in the experimental set-up (tubes, syringes etc.).
<ol>
	<li>Prepare the system as described in step 3, keeping electrodes and micropipette extended out of the guide tubes. A depth of at least 7,000 &#181;m is recommended to avoid loss of measurement volume due to adhesion along the outer surface of the micropipette and electrodes.</li>
	<li>Place the recording system in the position it will be used in during the experiment and put the syringe into the microinjection pump. Fix the syringe in place using the rubber band and adjustable grip (see Figure 1A). Slide the movable part of the pump until it is firmly in place behind the plunger of the syringe.</li>
	<li>Using the software-controlled motor unit, eject a volume large enough to be measured precisely, e.g., 1,000 nl. It is preferable to use one single step to eject the total volume in order to avoid effects of capillary action along the micropipette surface. Very low velocities (1 nl/s) can also lead to this effect during the validation procedure.</li>
	<li>Collect the total volume in a container placed under the micropipette, or carefully collect the ejected drop directly from the tip of the micropipette. Estimate the ejected volume using a pipette or by weighing with a precision scale.</li>
	<li>Repeat the procedure several times to confirm measurements.</li>
</ol>
5. Acute Recordings
<ol>
	<li>Set the x-y position of the recording system. This defines the point at which the guide tubes reach the dura mater within the chronically implanted recording chamber. Make sure the guide tubes are retracted completely (guide tube z position 0).</li>
	<li>Bring the recording system into position and place the syringe in the microinjection pump. Fix the syringe in place using the rubber band and adjustable grip (see Figure 1A). Slide the movable part of the pump until it is firmly in place behind the plunger of the syringe. If a drop of substance is visible at the tip of the guide tubes, carefully remove it using a sterile cotton bud.</li>
	<li>Prepare the animal for recording according to the laboratory&#39;s procedure (see18 for example guidelines).</li>
	<li>Securely mount the recording system on the recording chamber of the monkey.</li>
	<li>Slowly manually lower the guide tubes into the recording chamber until the dura is reached, then drive the electrodes using the motor control software.</li>
	<li>As it is not possible to measure impedance of the micropipette, first drive with electrodes and check their impedances regularly at different depths. After a penetration of the dura is successfully performed without damaging the electrodes, advance the micropipette.</li>
	<li>Drive the electrodes and the micropipette to the target electrode depth at which the brain area of interest is expected to be found. Slowly advance the electrode until it is close enough to record the activity of a single unit, as evidenced by a good signal-to-noise ratio in the recorded signal. Importantly, position the recording electrode and the micropipette at the same depth to ensure the minimum distance between electrode and micropipette.
	<ol>
		<li>If possible, keep the electrodes and micropipette at this depth for the entire recording. However, if the only way to maintain signal quality of the recorded cell is to move the electrodes, then drive the electrodes and the micropipette simultaneously to maintain the distance between them.</li>
	</ol>
	</li>
</ol>
6. Spatial Attention Task
<ol>
	<li>In a series of trials, present two moving dot patterns on the screen, one positioned within the receptive field of the recorded neuron and the other outside of it, together with a centrally presented fixation point that the animal has to foveate throughout each trial19,20. 
	Note: The monkey is trained to respond to a direction change in the cued dot pattern (the target event) while ignoring any direction change in the other dot pattern, and is rewarded with a drop of liquid for every successful completion of a trial19,20. As a sensory control condition, the monkey has to report a luminance change of the fixation point while ignoring both moving dot patterns (see Figure 2 for a more detailed description of the task).</li>
</ol>
7. Pharmacological Manipulation While Recording
Note: While the monkey is performing the task, inject the substance in a block-wise manner. Three consecutive blocks are defined: control, which acts as a baseline; injection, during which a substance is ejected; and recovery, during which the cells targeted by the injection return to baseline.
<ol>
	<li>During an injection block, inject a predefined amount of the substance at regular intervals e.g., 2 nl every minute at a rate of 2 nl/s. For this example, use scopolamine hydrochloride. The injection process is controlled using software which provides various options. For example, use the step function to define injection volume, and press the injection button every minute according to the clock of the recording software. 
	Note: The exact duration of the injection block is substance and experiment dependent, e.g., for scopolamine use 2 nl injections each minute for 10 min (20 nl in total). It is preferable not to advance the electrodes and micropipette during the injection block.</li>
	<li>Note the time and the trial during which the substance is injected, the depth of the electrodes and micropipette, as well as the amount of ejected substance.</li>
	<li>Follow the injection block with a recovery block, in which no substance is injected. The duration of the recovery block is substance specific and needs to be defined in pre-tests. Monitor and maintain the recording quality of the selected single units until the end of the recovery block.</li>
	<li>Repeat the three blocks for as long as the recording quality and motivation of the monkey allow.</li>
</ol>
8. Post Recording Procedures
<ol>
	<li>After data recording, retract electrodes and micropipette into the guide tubes and then manually retract the guide tubes. Remove the recording system from the recording chamber of the monkey. Release the syringe from the injection pump and transfer the system to the preparation area for cleaning.</li>
	<li>Handle the animal (including cleaning of the recording chamber18) according to the standard procedures of the laboratory and return it to the housing facility.</li>
	<li>Rinse the outside of the guide tubes with hydrogen peroxide (3%) and then with deionized water. Drive electrodes and micropipette out of the guide tubes, rinse with hydrogen peroxide and then deionized water.</li>
	<li>Exchange the barrel of the syringe with a barrel of a syringe filled with sterile saline, keeping the needle in the tube. Flush the tube and the micropipette with 1-2 ml of saline. After flushing, remove the barrel and fill it with air. Reinsert the barrel into the needle and dry the tube and the micropipette from the inside by gently pushing air through.</li>
	<li>Store the guide tubes, extended electrodes and micropipette immersed in the enzyme solution to avoid drying as well as to ensure the breakdown of organic material.</li>
</ol>

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The authors have nothing to disclose.

Here we have illustrated in detail how to perform reliable and precise injections and high quality single-cell recordings with an &#34;off-the-shelf&#34; pressure injection system. While this method of drug delivery has previously been used in behaving monkeys (reviewed in17), the system presented here has advantages, reviewed below.
As illustrated in Figure 4A, the system described here can provide stable measurement of single neuron activity with and without pharmacological injections in the direct vicinity of the recording site. As shown in Figure 4B, the injection of a control substance, saline, did not lead to a change in firing rate. This control demonstrates that the injection process itself has no measurable influence on the firing properties of the recorded neurons.
The spatial configuration of neuron, recording electrode, and micropipette is of crucial importance in these experiments. Although a precise measurement of their relative positions in the tissue during recording is not possible, we can consider and control for possible sources of variance. First, during volume injection there is a risk that the neuron of interest may be displaced away from the recording electrode, affecting the stability of recorded signals. For that reason it is prudent to compare the firing rate before and after the injection block to verify signal stability. Second, the guide tube configuration of the recording system defines the distance between electrodes and micropipette (e.g., 305 &#181;m in the concentric 3-channel system used in this experiment). As the system provides precise position control for the depth of electrodes and micropipette in the tissue, the distance between them can be minimized by carefully calibrating relative depth before recording (step 3.5), and keeping them at a common depth during recordings.
Potential limitations 
In addition to in-house quality control by the manufacturer, the system needs to be validated under lab conditions, as different brands of tubes, syringes etc. can be used and could lead to differences in ejected volumes. Although the system can be used to inject very small volumes as in the experiment shown here, these are below the minimum volume that can be validated due to practical measurement limits in a normal laboratory environment. However, larger injection volumes can be used to infer the relation between the software-defined volume and the volume ejected by the hardware. If transparent tubes are used, an additional visual check of the injection process is possible by measuring the displacement of a visual marker.
Inserting the micropipette into the system is more demanding than electrode insertion, as the diameter of the micropipette is slightly larger and the material is more fragile. In addition, joining the tube to the pin of the micropipette is challenging as it entails a high risk of breaking the upper part of the micropipette. However, the lifetime of a successfully loaded micropipette is several months, even with daily use.
In practice, we have not yet encountered a blockage in the injection system during post-recording cleaning of the system. Nevertheless, no &#34;online&#34; check is possible, and there is a risk that a physical blockage (such as tissue at the micropipette tip) might prevent substance injection. It might therefore be advisable to analyze the data conservatively, such as including only those cells in further analysis that show significant changes in firing rates between control and injection blocks of the experiment.
Despite their small diameter, microelectrodes and pipettes will displace brain tissue and may cause some local tissue damage. This can be minimized by manually positioning the tip of the guide tubes just above the dura mater. The electrodes then penetrate the dura and their intactness is inferred by measuring their impedances online. Afterwards, the micropipette is inserted. When using this approach, regular removal of tissue above the dura is recommended to further reduce the risk of electrode or pipette breakage.
Comparison to alternative methods 
The system used here shows clear advantages compared to other pressure injection systems. One strong advantage is the diameter of the micropipette (approximately 100 &#181;m), which is half the size of other available probes17 and therefore minimizes neural tissue damage. In contrast to previous designs, the current system employs spatially separated recording electrode and micropipette. Although other systems provide a smaller distance between electrode and pipette, the system described here allows independent depth changes of electrodes and pipette, thus permitting variable relative distances within a recording session. Importantly, no compromise regarding recording quality needs to be made, as the injection system is an extension of an established recording device. While only one micropipette and thus one substance is used in this protocol, it is possible to inject several substances within one experimental procedure. To achieve this, several micropipettes can be threaded into separate guide tubes and connected to syringes mounted in individual injection pumps. Finally, controlling the system is easy, as only one computer program is needed to advance the electrodes and micropipette, and to perform the pressure injection during the experiment.
Comparing pressure injection to iontophoresis, there are relative advantages and disadvantages. For example, pressure injection requires greater volumes to be introduced into the tissue than iontophoresis, thus increasing the risk of neuronal displacement. The current protocol used volumes in the nL range, and we rarely experienced noticeable changes in a recorded cell&#39;s signal quality. The system also allows larger volumes to be injected, which is potentially useful for behavioral manipulations but could impact stability of neuronal recording. A clear advantage of pressure injection over iontophoresis is the larger variety of useable substances as there is no requirement to use charged substances. However, pH values should be checked and compared between experimental and control substances (e.g., saline).
The question might arise why to use the long-established method of pressure injection instead of newer techniques such as optogenetics for manipulating neural activity. Although well established in rodents, optogenetics is not yet reliably established in rhesus monkeys. In particular, it does not yet allow the local manipulation of cells selective for a particular neurotransmitter type. In the longer run, we see great potential for the combination of the advantages of pharmacological manipulations with the advantages of optogentic manipulations in elucidating the neural basis of cognitive functions.
Here we have shown how pressure injection can be used to pharmacologically manipulate a locally restricted area in the brain of awake, behaving rhesus monkeys. We hope that this method inspires other scientists to investigate neuromodulatory contributions to the dynamics of neuronal activity.

In cortical and subcortical areas, neuronal activity is affected by various neuromodulators, for example acetylcholine1. These modulatory effects on neuronal responses have been reported in in vitro studies2, as well as in electrophysiological recordings from anesthetized animals3 and systemic pharmacological manipulations in humans4. Nevertheless, the exact role of different neuromodulators and the involvement of various receptor subtypes are largely unknown. To measure the effects of specific neuromodulators on the activity of single neurons, it is desirable to induce a temporary neuromodulator change as close as possible to the recording electrode. Furthermore, it is important that those manipulations are done in awake animals, as cognitive functions are only present in the absence of anesthesia. Additionally, anesthesia interacts with cholinergic and GABAergic systems5,6 and can lead to changes in neural activity3.
Within the last decades, two main methods of local drug delivery have been developed and refined: iontophoresis and pressure injection. In both methods drugs are delivered through micropipettes made of either glass or steel. With iontophoresis, an electrical current regulates the release of the drug7. Additionally, there is a significant contribution of electro-osmosis to the total amount of ejected molecules8, correlating with the tip diameter9 of the micropipette as well as with the concentration8 of the substance used. Iontophoresis is a powerful tool to quickly and precisely manipulate small volumes of nervous tissue. For iontophoretic injections, multi-barrel micropipettes are usually used10, with one acting as a recording device while the other positions serve as delivery pipettes. A limitation of this method is that only charged molecules can be used, severely limiting the selection of drugs. 
Pressure injection uses either air compression or mechanical pressure to eject a substance from a micropipette. Using this method any soluble substance, charged or uncharged, can be used, including large molecules. The method of pressure injection was first described by Reyniers in 1933 and further refined in the 1950s (see Lalley11 for a review). In the 1980s the method was further refined to allow delivery of amounts in the nanoliter range (mainly lidocain12) to a defined brain area13 while simultaneously performing single-cell recording. The ejected volume was usually monitored by observing the movement of a marker, such as the meniscus in the upper part of the pipette13. Pressure injection was first used in the 1990s in awake animals, both extracellularly14 and intracellularly15,16. Based on the cumulative expertise gained in these studies it is now possible to reliably record from different brain structures in combination with pharmacological manipulation (see17 for a comparison of recent pressure injection systems). 
An enduring open issue for both drug delivery methods is the difficulty in determining the precise volume injected. This is an even bigger challenge for experiments with awake, behaving rhesus monkeys where the animal performs the experimental task in a separate room. This can be alleviated by the use of a software-controlled system instead of relying on a visual marker to continuously monitor an injection. 
The system described here is an extension of a well-established electrophysiological recording system (Mini Matrix System) and combines an injection pipette with multiple parallel-oriented recording electrodes at defined distances in a customizable arrangement. Pharmacological manipulation of the tissue near the recording electrode is possible using only a small amount of substance, ensuring a fast recovery and allowing multiple blocks of injection and control/recovery within the limited time window offered by the behavioral task of the animal.

<table><tbody><tr><td>(-)-Scopolamine hydrochloride</td><td>Sigma-Aldrich</td><td>55-16-3</td><td>Mr 339.81 g/mol</td></tr><tr><td>NaCl 0.9%&amp;nbsp;</td><td>B. Braun Melsungen AG</td><td>3079870</td><td>5ml&amp;nbsp;</td></tr><tr><td>Terg-a-zyme</td><td>Sigma-Aldrich</td><td>Z273287</td><td>enzyme detergen</td></tr><tr><td>Hydrogen peroxide</td><td>Roth</td><td></td><td>Used in 3% solution with deionized water</td></tr><tr><td>Ethanol</td><td>Chemie-Vertieb Hannover</td><td>104642</td><td>70%</td></tr><tr><td>Deionized water</td><td></td><td></td><td></td></tr><tr><td>Injekt 40 Duo</td><td>B. Braun Melsungen AG</td><td>9166432V</td><td>Syringe and needle&amp;nbsp;</td></tr><tr><td>Eppendorf Safe-Lock microcentrifuge tubes, amber</td><td>Eppendorf</td><td>0030 120.191</td><td>1,5ml&amp;nbsp;</td></tr><tr><td>Quarzglass micropipette</td><td>Thomas Recording</td><td></td><td></td></tr><tr><td>Recording electrode</td><td>Thomas Recording</td><td></td><td>quartz/platinum-tungsten fiber electrode; impedance value 1-2 M&amp;Omega;&amp;nbsp; and 0.3-0.5 M&amp;Omega;</td></tr><tr><td>PharmedBPT-Schlauch</td><td>Saint-Gobain Performance Plastics&amp;nbsp;</td><td>3702003</td><td>&amp;nbsp;Size: 0,25 x 2,05 mm (Wd: 0,9mm)</td></tr><tr><td>Loctite 401</td><td>Henkel</td><td>233641</td><td>Superglue</td></tr><tr><td>Silicon oil</td><td>Thomas Recording</td><td>M-1000</td><td></td></tr><tr><td>Minisart RC15</td><td>Sartorius</td><td>17761----------R</td><td>Syringe filter</td></tr><tr><td>Multichannel Micro Injection System</td><td>Thomas Recording</td><td></td><td>multichannel microelectrode manipulator &amp;ldquo;System Eckhorn&amp;rdquo; equipped with microelectrodes and micropipettes and a precision multichannel microinjection pump</td></tr><tr><td>McLab</td><td>custom</td><td></td><td>internal lab software to control stimulus presentation</td></tr></tbody></table>

a pressure injection system for investigating the neuropharmacology of information processing in awake behaving macaque monkey cortex

The top-down modulation of feed-forward cortical information processing is functionally important for many cognitive processes, including the modulation of sensory information processing by attention. However, little is known about which neurotransmitter systems are involved in such modulations. A practical way to address this question is to combine single-cell recording with local and temporary neuropharmacological manipulation in a suitable animal model. Here we demonstrate a technique combining acute single-cell recordings with the injection of neuropharmacological agents in the direct vicinity of the recording electrode. The video shows the preparation of the pressure injection/recording system, including preparation of the substance to be injected. We show a rhesus monkey performing a visual attention task and the procedure of single-unit recording with block-wise pharmacological manipulations.

Figure 2 depicts the spatial attention task the monkey performed while the injection process was conducted. The monkey was trained to attend to either the stimulus located within the receptive field of the recorded neuron (attend-in), the stimulus located outside of the receptive field (attend-out) or the fixation point (attend-fix). These conditions allow a comparison of neuronal activity in different attentional states.
Figure 3 shows a peri-stimulus time histogram of a sample neuron in an experiment using scopolamine, a muscarinic cholinergic antagonist. The plot demonstrates the response suppression during scopolamine injection versus no injection, when a pattern moving in the cell&#39;s preferred direction is presented inside the neuron&#39;s receptive field and is attended by the animal. The two first peaks represent the neuron&#39;s response to the on- and offset of the spatial cue, which appears inside its receptive field. This is followed by the response to the moving pattern which appears on the screen 500 ms after cue onset. The gray shaded area depicts the analysis period used to calculate the average firing rate for every trial. The green area highlights the suppressive influence of scopolamine injection on the cell&#39;s firing rate. The dark green region shows the suppression within the analysis period.
Figure 4A shows the effect of scopolamine on the average firing rate of the sample neuron in each of the three attentional conditions. The neuron&#39;s firing rate for the two spatial attention conditions (attention inside or outside of the receptive field of the recording neuron) as well as for the sensory condition (attention at fixation point) dropped shortly after the first injection of the injection block (grey shaded area) and during the recovery block increased after a delay to the same level as before the injection.
Figure 4B shows a control recording from a second sample neuron in which saline (0.9% NaCl) was injected, using the same protocol as for the scopolamine injection. During the injection block no change in the neuron&#39;s firing rate was observed compared to the control block.
<img alt="Figure 1" src="/files/ftp_upload/53724/53724fig1.jpg" /> 
Figure 1. Set-up used for pharmacological manipulation while recording. (A) Depicts the microinjection pump and the electrophysiological recording system equipped with electrodes and micropipette. The guidetube gap, into which silicon oil is inserted to lubricate the electrodes and micropipette, is shown enlarged. (B) Displays an example micropipette (above) and recording electrode (below). For size comparison, a euro cent (diameter: 16 mm) is placed underneath. <a href="https://www.jove.com/files/ftp_upload/53724/53724fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53724/53724fig2.jpg" /> 
Figure 2. Task design to guide spatial attention. Monkeys were trained to detect a motion direction change in the cued dot pattern. The cue was either placed within the neuron&#39;s receptive field (attend-in), as shown in the figure, or outside of it (attend-out). As a sensory control, the monkey was trained to detect a luminance change of the fixation point (attend-fix). <a href="https://www.jove.com/files/ftp_upload/53724/53724fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53724/53724fig3.jpg" /> 
Figure 3. Influence of antagonist scopolamine on firing rate. The peri-stimulus time histogram for a sample neuron is shown for the attend-in condition (attention inside the receptive field of the recorded neuron) during the injection block and during the control block. The x-axis depicts the time in milliseconds after cue onset and the y-axis shows the firing rate in spikes/sec. The grey area depicts the analysis period (300-800 ms after stimulus onset) used to calculate the trial-averaged firing rate. The green shaded area shows the suppression in firing rate across the two conditions. The dark green color highlights the suppression within the analysis period. <a href="https://www.jove.com/files/ftp_upload/53724/53724fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53724/53724fig4.jpg" /> 
Figure 4. Effect of scopolamine and saline on firing rate. (A) Antagonist scopolamine injection. The trial-averaged firing rate of the sample cell from Figure 3 over the course of the experiment is shown for the preferred stimulus for all three attentional conditions. The x-axis depicts the trial start time in minutes and the y-axis shows the unit&#39;s firing rate in spikes per seconds. Symbols (<img alt="Plus" src="/files/ftp_upload/53724/53724plussign.jpg" /> attend-in, <img alt="Circle" src="/files/ftp_upload/53724/53724circle.jpg" /> attend-fix, <img alt="Trianle" src="/files/ftp_upload/53724/53724triangle.jpg" /> attend-out) represent the neuron&#39;s firing rate within the analysis period in every successfully performed trial, and horizontal lines (solid line: attend-in, dotted line: attend-fix, dashed line: attend-out) show average firing rate for the three different experimental blocks (control, injection, recovery). The grey shaded area shows the injection block, beginning with the first injection and ending 1 min after the last injection. During the injection block 2 nL of 0.1 molar scopolamine were injected every minute with an injection velocity of 2 nl/s. (B) Saline injection. The firing rate of a sample cell over the course of the control experiment is shown for the preferred stimulus for all three attentional conditions. Grey shaded area visualizes the block of saline injection. <a href="https://www.jove.com/files/ftp_upload/53724/53724fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Pressure Injection System for Investigating the Neuropharmacology of Information Processing in Awake Behaving Macaque Monkey Cortex at JoVE.com

A Pressure Injection System for Investigating the Neuropharmacology of Information Processing in Awake Behaving Macaque Monkey Cortex

Here, we show the pressure injection of neuropharmacological substances during single-cell recording in an awake, behaving macaque monkey. This procedure allows pharmacological manipulation in the direct vicinity of a cortical recording site.

The top-down modulation of feed-forward cortical information processing is functionally important for many cognitive processes, including the modulation ...

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8.4K Views.  German Primate Center. Here, we show the pressure injection of neuropharmacological substances during single-cell recording in an awake, behaving macaque monkey. This procedure allows pharmacological manipulation in the direct vicinity of a cortical recording site. 

Video: A Pressure Injection System for Investigating the Neuropharmacology of Information Processing in Awake Behaving Macaque Monkey Cortex

Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

It is well known that anesthesia alters neural response properties in various regions of the brain.13. In the auditory system, fundamental response properties of brainstem neurons including threshold, frequency specificity, and inhibitory sidebands are altered in significant ways under anesthesia1-2. These observations prompted physiologists to seek ways to record from single neurons without the contaminating effects of anesthesia. One result was a decerebrate preparation, where the brainstem was completely transected at the level of the midbrain4. The drawbacks of this preparation are a formidable surgery, the elimination of descending projections from the forebrain, and an inability to use sensory stimulation to examine structures above the midbrain. A different strategy has been to implant electrode arrays chronically to record from single neurons and multiunit clusters while the animal is awake and/or behaving5,6. These techniques however are not compatible with injecting tracer dyes after first electrophysiologically characterizing a brain structure. To avoid altering neural response properties with anesthetics while recording electrophysiological response properties from single neurons, we have adapted a head restraint technique long used in bats7-9 to mouse10-12. Using this method, we are able to conduct electrophysiological recordings over several days in the unanesthetized mouse. At the end of the recording sessions, we can then inject a dye to reconstruct electrode positions and recording sites or inject a tracer so that pathways to and from the recording loci can be determined. This method allows for well isolated single neuron recordings over multiple days without the use anesthetics.

Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

It is well known that anesthesia alters neural response properties in various regions of the brain.13.  In the auditory system, fundamental response ...

Neuroscience

Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, direction5-7, ocular dominance8,9 and binocular disparity9. Orientation selectivity is the most widely studied feature and a continuous map with a quasi-periodic layout for preferred orientation is present across the entire primary visual cortex10,11. Integrating the synaptic, cellular and network contributions that lead to stimulus selective responses in these functional maps requires the hybridization of imaging techniques that span sub-micron to millimeter spatial scales. With conventional intrinsic signal optical imaging, the overall layout of functional maps across the entire surface of the visual cortex can be determined12. The development of in vivo two-photon microscopy using calcium sensitive dyes enables one to determine the synaptic input arriving at individual dendritic spines13 or record activity simultaneously from hundreds of individual neuronal cell bodies6,14. Consequently, combining intrinsic signal imaging with the sub-micron spatial resolution of two-photon microscopy offers the possibility of determining exactly which dendritic segments and cells contribute to the micro-domain of any functional map in the neocortex. Here we demonstrate a high-yield method for rapidly obtaining a cortical orientation map and targeting a specific micro-domain in this functional map for labeling neurons with fluorescent dyes in a non-rodent mammal. With the same microscope used for two-photon imaging, we first generate an orientation map using intrinsic signal optical imaging. Then we show how to target a micro-domain of interest using a micropipette loaded with dye to either label a population of neuronal cell bodies or label a single neuron such that dendrites, spines and axons are visible in vivo. Our refinements over previous methods facilitate an examination of neuronal structure-function relationships with sub-cellular resolution in the framework of neocortical functional architectures.

A method is described for labeling neurons with fluorescent dyes in predetermined functional micro-domains of the neocortex. First, intrinsic signal optical imaging is used to obtain a functional map. Then two-photon microscopy is used to label and image neurons within a micro-domain of the map.

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, ...

Juxtacellular Monitoring and Localization of Single Neurons within Sub-cortical Brain Structures of Alert, Head-restrained Rats

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures in behaving animals remains a technical challenge, especially if the structures must be targeted stereotaxically. This protocol describes convenient surgical and electrophysiological techniques that maintain the animal&#8217;s head in the stereotaxic plane and unambiguously isolate the spiking activity of single neurons. The protocol combines head restraint of alert rodents, juxtacellular monitoring with micropipette electrodes, and iontophoretic dye injection to identify the neuron location in post-hoc histology. While each of these techniques is in itself well-established, the protocol focuses on the specifics of their combined use in a single experiment. These neurophysiological and neuroanatomical techniques are combined with behavioral monitoring. In the present example, the combined techniques are used to determine how self-generated vibrissa movements are encoded in the activity of neurons within the somatosensory thalamus. More generally, it is straightforward to adapt this protocol to monitor neuronal activity in conjunction with a variety of behavioral tasks in rats, mice, and other animals. Critically, the combination of these methods allows the experimenter to directly relate anatomically-identified neurophysiological signals to behavior.

This protocol describes the design and surgical implantation of a head-restraining mechanism to monitor neuronal activity in sub-cortical brain structures in alert rats. It delineates procedures to isolate single neurons in the juxtacellular configuration and to efficiently identify their anatomical locations.

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Induction of an Isoelectric Brain State to Investigate the Impact of Endogenous Synaptic Activity on Neuronal Excitability In Vivo

The way neurons process information depends both on their intrinsic membrane properties and on the dynamics of the afferent synaptic network. In particular, endogenously-generated network activity, which strongly varies as a function of the state of vigilance, significantly modulates neuronal computation. To investigate how different spontaneous cerebral dynamics impact single neurons&#39; integrative properties, we developed a new experimental strategy in the rat consisting in suppressing in vivo all cerebral activity by means of a systemic injection of a high dose of sodium pentobarbital. Cortical activities, continuously monitored by combined electrocorticogram (ECoG) and intracellular recordings are progressively slowed down, leading to a steady isoelectric profile. This extreme brain state, putting the rat into a deep comatose, was carefully monitored by measuring the physiological constants of the animal throughout the experiments. Intracellular recordings allowed us to characterize and compare the integrative properties of the same neuron embedded into physiologically relevant cortical dynamics, such as those encountered in the sleep-wake cycle, and when the brain was fully silent.

Induction of an Isoelectric Brain State to Investigate the Impact of Endogenous Synaptic Activity on Neuronal Excitability In Vivo

This procedure performs long-lasting in vivo intracellular recordings from single neurons during physiologically relevant cerebral states and after complete abolition of ongoing electrical activities, resulting in an isoelectric brain state. The physiological constants of the animal are carefully monitored during the transition to the artificial comatose condition.

The way neurons process information depends both on their intrinsic membrane properties and on the dynamics of the afferent synaptic network. In ...

Convection Enhanced Delivery of Optogenetic Adeno-associated Viral Vector to the Cortex of Rhesus Macaque Under Guidance of Online MRI Images

In non-human primate (NHP) optogenetics, infecting large cortical areas with viral vectors is often a difficult and time-consuming task. Here, we demonstrate the use of magnetic resonance (MR)-guided convection enhanced delivery (CED) of optogenetic viral vectors into primary somatosensory (S1) and motor (M1) cortices of macaques to obtain efficient, widespread cortical expression of light-sensitive ion channels. Adeno-associated viral (AAV) vectors encoding the red-shifted opsin C1V1 fused to yellow fluorescent protein (EYFP) were injected into the cortex of rhesus macaques under MR-guided CED. Three months post-infusion, epifluorescent imaging confirmed large regions of optogenetic expression (&#62;130 mm2) in M1 and S1 in two macaques. Furthermore, we were able to record reliable light-evoked electrophysiology responses from the expressing areas using micro-electrocorticographic arrays. Later histological analysis and immunostaining against the reporter revealed widespread and dense optogenetic expression in M1 and S1 corresponding to the distribution indicated by epifluorescent imaging. This technique enables us to obtain expression across large areas of the cortex within a shorter period of time with minimal damage compared to the traditional techniques and can be an optimal approach for optogenetic viral delivery in large animals such as NHPs. This approach demonstrates great potential for network-level manipulation of neural circuits with cell-type specificity in animal models evolutionarily close to humans.&#160;

Here, we demonstrate magnetic resonance (MR)-guided convection enhanced delivery (CED) of viral vectors into the cortex as an efficient and simplified approach for achieving optogenetic expression across large cortical areas in the macaque brain.

In non-human primate (NHP) optogenetics, infecting large cortical areas with viral vectors is often a difficult and time-consuming task. Here, we ...

Microinjectrode System for Combined Drug Infusion and Electrophysiology

This microinjectrode system is designed for drug infusion, electrophysiology, and delivery and retrieval of experimental probes, such as microelectrodes and nanosensors, optimized for repeated use in awake, behaving animals. The microinjectrode system can be configured for multiple purposes: (1) simple arrangement of the cannula for placement of an experimental probe that would otherwise be too fragile to penetrate the dura mater, (2) microfluidic infusion of a drug, either independently or coupled to a cannula containing an experimental probe (i.e., microelectrode, nanosensor). In this protocol we explain the step by step construction of the microinjectrode, its coupling to microfluidic components, and the protocol for use of the system in vivo. The microfluidic components of this system allow for delivery of volumes on the nanoliter scale, with minimal penetration damage. Drug infusion can be performed independently or simultaneously with experimental probes such as microelectrodes or nanosensors in an awake, behaving animal. Applications of this system range from measuring the effects of a drug on cortical electrical activity and behavior, to understanding the function of a specific region of cortex in the context of behavioral performance based on probe or nanosensor measurements. To demonstrate some of the capabilities of this system, we present an example of muscimol infusion for reversible inactivation of the frontal eye field (FEF) in rhesus macaque during a working memory task.

We present a microinjectrode system designed for electrophysiology and assisted delivery of experimental probes (i.e., nanosensors, microelectrodes), with optional drug infusion. Widely available microfluidic components are coupled to a cannula containing the probe. A step-by-step protocol for microinjectrode construction is included, with results during muscimol infusion in macaque cortex.

This microinjectrode system is designed for drug infusion, electrophysiology, and delivery and retrieval of experimental probes, such as microelectrodes ...

Injections of AAV Vectors for Optogenetics in Anesthetized and Awake Behaving Non-Human Primate Brain

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of optogenetics, however, requires that safety and efficacy be demonstrated in animal models, ideally in non-human primates (NHPs), because of their neurological similarity to humans. The number of candidate vectors that are potentially useful for neuroscience and medicine is vast, and no high-throughput means to test these vectors yet exists. Thus, there is a need for techniques to make multiple spatially and volumetrically accurate injections of viral vectors into NHP brain that can be identified unambiguously through postmortem histology. Described herein is such a method. Injection cannulas are constructed from coupled polytetrafluoroethylene and stainless-steel tubes. These cannulas are autoclavable, disposable, and have low minimal-loading volumes, making them ideal for the injection of expensive, highly concentrated viral vector solutions. An inert, red-dyed mineral oil fills the dead space and forms a visible meniscus with the vector solution, allowing instantaneous and accurate measurement of injection rates and volumes. The oil is loaded into the rear of the cannula, reducing the risk of co-injection with the vector. Cannulas can be loaded in 10 min, and injections can be made in 20 min. This procedure is well suited for injections into awake or anesthetized animals. When used to deliver high-quality viral vectors, this procedure can produce robust expression of optogenetic proteins, allowing optical control of neural activity and behavior in NHPs.

As currently implemented, optogenetics in non-human primates requires injection of viral vectors into the brain. An optimal injection method should be reliable and, for many applications, capable of targeting individual sites of arbitrary depth that are readily and unambiguously identified in postmortem histology. An injection method with these properties is presented.

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of ...

Head Implants for the Neuroimaging of Awake, Head-Fixed Rats

Anesthetics, commonly used in preclinical and fundamental scientific research, have a depressive influence on the metabolic, neuronal, and vascular functions of the brain and can adversely influence neurophysiological results. The use of awake animals for research studies is advantageous but poses the major challenge of keeping the animals calm and stationary to minimize motion artifacts throughout data acquisition. Awake imaging in smaller-sized rodents (e.g., mice) is very common but remains scant in rats as rats are bigger, stronger, and have a greater tendency to oppose movement restraints and head fixation over the long durations required for imaging. A new model of neuroimaging of awake, head-fixed rats using customized hand-sewn slings, 3D-printed head implants, head caps, and a headframe is described. The results acquired following a single trial of single-whisker stimulation suggest an increase in the intensity of the evoked functional response. The acquisition of the evoked functional response from awake, head-fixed rats is faster than that from anesthetized rats, reliable, reproducible, and can be used for repeated longitudinal studies.

A detailed new procedure for functional imaging of awake, head-fixed rats is described.

Anesthetics, commonly used in preclinical and fundamental scientific research, have a depressive influence on the metabolic, neuronal, and vascular ...

Drug Microinjection with Electrophysiological Recording in a Rat Brain

Source: Lai, J., et al. <a href="https://app.jove.com/v/52271">Simultaneous electrophysiological recording and micro-injections of inhibitory agents in the rodent brain.</a> J. Vis. Exp. (2015)
This protocol outlines the assembly and use of an injectrode for precise drug delivery into targeted brain regions of an anesthetized rat, followed by monitoring of neuronal activity through recorded electrical signals.

A Pressure Injection System for Investigating the Neuropharmacology of Information Processing in Awake Behaving Macaque Monkey Cortex

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