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 border="0" cellpadding="0" cellspacing="0" width="560">
	<tbody>
		<tr height="41">
			<td height="41" style="height:41px;width:141px;">(-)-Scopolamine hydrochloride</td>
			<td style="width:88px;">Sigma-Aldrich</td>
			<td style="width:87px;">55-16-3</td>
			<td style="width:244px;">Mr 339.81 g/mol</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:141px;">NaCl 0.9%&#160;</td>
			<td style="width:88px;">B. Braun Melsungen AG</td>
			<td style="width:87px;">3079870</td>
			<td style="width:244px;">5ml&#160;</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:141px;">Terg-a-zyme</td>
			<td style="width:88px;">Sigma-Aldrich</td>
			<td style="width:87px;">Z273287</td>
			<td style="width:244px;">enzyme detergen</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:141px;">Hydrogen peroxide</td>
			<td style="width:88px;">Roth</td>
			<td style="width:87px;"></td>
			<td style="width:244px;">Used in 3% solution with deionized water</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:141px;">Ethanol</td>
			<td style="width:88px;">Chemie-Vertieb Hannover</td>
			<td style="width:87px;">104642</td>
			<td style="width:244px;">70%</td>
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		<tr height="22">
			<td height="22" style="height:22px;width:141px;">Deionized water</td>
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			<td style="width:244px;"></td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:141px;">Injekt 40 Duo</td>
			<td style="width:88px;">B. Braun Melsungen AG</td>
			<td style="width:87px;">9166432V</td>
			<td style="width:244px;">Syringe and needle&#160;</td>
		</tr>
		<tr height="98">
			<td height="120" rowspan="2" style="height:121px;width:141px;">Eppendorf Safe-Lock microcentrifuge tubes, amber</td>
			<td rowspan="2" style="width:88px;">Eppendorf</td>
			<td rowspan="2" style="width:87px;">0030 120.191</td>
			<td rowspan="2" style="width:244px;">1,5ml&#160;</td>
		</tr>
		<tr height="22">
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:141px;">Quarzglass micropipette</td>
			<td style="width:88px;">Thomas Recording</td>
			<td style="width:87px;"></td>
			<td style="width:244px;"></td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:141px;">Recording electrode</td>
			<td style="width:88px;">Thomas Recording</td>
			<td style="width:87px;"></td>
			<td style="width:244px;">quartz/platinum-tungsten fiber electrode; impedance value 1-2 M&#937;&#160; and 0.3-0.5 M&#937;</td>
		</tr>
		<tr height="38">
			<td height="81" rowspan="3" style="height:82px;width:141px;">PharmedBPT-Schlauch</td>
			<td rowspan="3" style="width:88px;">Saint-Gobain Performance Plastics&#160;</td>
			<td rowspan="3" style="width:87px;">3702003</td>
			<td rowspan="3" style="width:244px;">&#160;Size: 0,25 x 2,05 mm (Wd: 0,9mm)</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="22">
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:141px;">Loctite 401</td>
			<td style="width:88px;">Henkel</td>
			<td style="width:87px;">233641</td>
			<td style="width:244px;">Superglue</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:141px;">Silicon oil</td>
			<td style="width:88px;">Thomas Recording</td>
			<td style="width:87px;">M-1000</td>
			<td style="width:244px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:141px;">Minisart RC15</td>
			<td style="width:88px;">Sartorius</td>
			<td style="width:87px;">17761----------R</td>
			<td style="width:244px;">Syringe filter</td>
		</tr>
		<tr height="121">
			<td height="121" style="height:121px;width:141px;">Multichannel Micro Injection System</td>
			<td style="width:88px;">Thomas Recording</td>
			<td style="width:87px;"></td>
			<td style="width:244px;">multichannel microelectrode manipulator &#8220;System Eckhorn&#8221; equipped with microelectrodes and micropipettes and a precision multichannel microinjection pump</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:141px;">McLab</td>
			<td style="width:88px;">custom</td>
			<td style="width:87px;"></td>
			<td style="width:244px;">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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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 ...

a-pressure-injection-system-for-investigating-neuropharmacology

Nanyang Technological University

Research

JoVE Journal

Behavior

8.3K 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

Construction of Microdrive Arrays for Chronic Neural Recordings in Awake Behaving Mice

State-of-the-art electrophysiological recordings from the brains of freely behaving animals allow researchers to simultaneously examine local field potentials (LFPs) from populations of neurons and action potentials from individual cells, as the animal engages in experimentally relevant tasks. Chronically implanted microdrives allow for brain recordings to last over periods of several weeks. Miniaturized drives and lightweight components allow for these long-term recordings to occur in small mammals, such as mice. By using tetrodes, which consist of tightly braided bundles of four electrodes in which each wire has a diameter of 12.5 &mu;m, it is possible to isolate physiologically active neurons in superficial brain regions such as the cerebral cortex, dorsal hippocampus, and subiculum, as well as deeper regions such as the striatum and the amygdala. Moreover, this technique insures stable, high-fidelity neural recordings as the animal is challenged with a variety of behavioral tasks. This manuscript describes several techniques that have been optimized to record from the mouse brain. First, we show how to fabricate tetrodes, load them into driveable tubes, and gold-plate their tips in order to reduce their impedance from M&Omega; to K&Omega; range. Second, we show how to construct a custom microdrive assembly for carrying and moving the tetrodes vertically, with the use of inexpensive materials. Third, we show the steps for assembling a commercially available microdrive (Neuralynx VersaDrive) that is designed to carry independently movable tetrodes. Finally, we present representative results of local field potentials and single-unit signals obtained in the dorsal subiculum of mice. These techniques can be easily modified to accommodate different types of electrode arrays and recording schemes in the mouse brain.

The design and assembly of microdrives for in vivo electrophysiological recordings of brain signals from the mouse is described. By attaching microelectrode bundles to sturdy driveable carriers, these techniques allow for long-term and stable neural recordings. The lightweight design allows for unrestricted behavioral performance by the animal following drive implantation.

State-of-the-art electrophysiological recordings from the brains of freely behaving animals allow researchers to simultaneously examine local field ...

Assessment of Murine Exercise Endurance Without the Use of a Shock Grid: An Alternative to Forced Exercise

Using laboratory mouse models, the molecular pathways responsible for the metabolic benefits of endurance exercise are beginning to be defined. The most common method for assessing exercise endurance in mice utilizes forced running on a motorized treadmill equipped with a shock grid. Animals who quit running are pushed by the moving treadmill belt onto a grid that delivers an electric foot shock; to escape the negative stimulus, the mice return to running on the belt. However, avoidance behavior and psychological stress due to use of a shock apparatus can interfere with quantitation of running endurance, as well as confound measurements of postexercise serum hormone and cytokine levels. Here, we demonstrate and validate a refined method to measure running endurance in na&#239;ve C57BL/6 laboratory mice on a motorized treadmill without utilizing a shock grid. When mice are preacclimated to the treadmill, they run voluntarily with gait speeds specific to each mouse. Use of the shock grid is replaced by gentle encouragement by a human operator using a tongue depressor, coupled with sensitivity to the voluntary willingness to run on the part of the mouse. Clear endpoints for quantifying running time-to-exhaustion for each mouse are defined and reflected in behavioral signs of exhaustion such as splayed posture and labored breathing. This method is a humane refinement which also decreases the confounding effects of stress on experimental parameters.

A method to assess exercise endurance in laboratory mice without the use of a shock grid is demonstrated. This method is a humane refinement that can decrease the confounding effects of stress on experimental parameters.

Using laboratory mouse models, the molecular pathways responsible for the metabolic benefits of endurance exercise are beginning to be defined. The most ...

Infant Auditory Processing and Event-related Brain Oscillations

Rapid auditory processing and acoustic change detection abilities play a critical role in allowing human infants to efficiently process the fine spectral and temporal changes that are characteristic of human language. These abilities lay the foundation for effective language acquisition; allowing infants to hone in on the sounds of their native language. Invasive procedures in animals and scalp-recorded potentials from human adults suggest that simultaneous, rhythmic activity (oscillations) between and within brain regions are fundamental to sensory development; determining the resolution with which incoming stimuli are parsed. At this time, little is known about oscillatory dynamics in human infant development. However, animal neurophysiology and adult EEG data provide the basis for a strong hypothesis that rapid auditory processing in infants is mediated by oscillatory synchrony in discrete frequency bands. In order to investigate this, 128-channel, high-density EEG responses of 4-month old infants to frequency change in tone pairs, presented in two rate conditions (Rapid: 70 msec ISI and Control: 300 msec ISI) were examined. To determine the frequency band and magnitude of activity, auditory evoked response averages were first co-registered with age-appropriate brain templates. Next, the principal components of the response were identified and localized using a two-dipole model of brain activity. Single-trial analysis of oscillatory power showed a robust index of frequency change processing in bursts of Theta band (3 - 8 Hz) activity in both right and left auditory cortices, with left activation more prominent in the Rapid condition. These methods have produced data that are not only some of the first reported evoked oscillations analyses in infants, but are also, importantly, the product of a well-established method of recording and analyzing clean, meticulously collected, infant EEG and ERPs. In this article, we describe our method for infant EEG net application, recording, dynamic brain response analysis, and representative results.

High-density electroencephalography (dEEG) is being used increasingly to study brain development and plasticity in the early years of life. Here we present an application of sophisticated analysis techniques that builds on traditional EEG recording to understand the oscillatory dynamics of rapid auditory processing in the infant brain.

Rapid auditory processing and acoustic change detection abilities play a critical role in allowing human infants to efficiently process the fine spectral ...

A Novel Experimental and Analytical Approach to the Multimodal Neural Decoding of Intent During Social Interaction in Freely-behaving Human Infants

Understanding typical and atypical development remains one of the fundamental questions in developmental human neuroscience. Traditionally, experimental paradigms and analysis tools have been limited to constrained laboratory tasks and contexts due to technical limitations imposed by the available set of measuring and analysis techniques and the age of the subjects. These limitations severely limit the study of developmental neural dynamics and associated neural networks engaged in cognition, perception and action in infants performing &#8220;in action and in context&#8221;. This protocol presents a novel approach to study infants and young children as they freely organize their own behavior, and its consequences in a complex, partly unpredictable and highly dynamic environment. The proposed methodology integrates synchronized high-density active scalp electroencephalography (EEG), inertial measurement units (IMUs), video recording and behavioral analysis to capture brain activity and movement non-invasively in freely-behaving infants. This setup allows for the study of neural network dynamics in the developing brain, in action and context, as these networks are recruited during goal-oriented, exploration and social interaction tasks.

This protocol presents a novel methodology for the neural decoding of intent from freely-behaving infants during unscripted social interaction with an actor. Neural activity is acquired using non-invasive high-density active scalp electroencephalography (EEG). Kinematic data is collected with inertial measurement units and supplemented with synchronized video recording.

Understanding typical and atypical development remains one of the fundamental questions in developmental human neuroscience. Traditionally, experimental ...

A Method to Quantify Visual Information Processing in Children Using Eye Tracking

Visual problems that occur early in life can have major impact on a child's development. Without verbal communication and only based on observational methods, it is difficult to make a quantitative assessment of a child's visual problems. This limits accurate diagnostics in children under the age of 4 years and in children with intellectual disabilities. Here we describe a quantitative method that overcomes these problems. The method uses a remote eye tracker and a four choice preferential looking paradigm to measure eye movement responses to different visual stimuli. The child sits without head support in front of a monitor with integrated infrared cameras. In one of four monitor quadrants a visual stimulus is presented. Each stimulus has a specific visual modality with respect to the background, e.g., form, motion, contrast or color. From the reflexive eye movement responses to these specific visual modalities, output parameters such as reaction times, fixation accuracy and fixation duration are calculated to quantify a child's viewing behavior. With this approach, the quality of visual information processing can be assessed without the use of communication. By comparing results with reference values obtained in typically developing children from 0-12 years, the method provides a characterization of visual information processing in visually impaired children. The quantitative information provided by this method can be advantageous for the field of clinical visual assessment and rehabilitation in multiple ways. The parameter values provide a good basis to: (i) characterize early visual capacities and consequently to enable early interventions; (ii) compare risk groups and follow visual development over time; and (iii), construct an individual visual profile for each child.

A method is described to quantify the quality of visual information processing based on reflexive eye movements in response to specific visual modalities. Reaction times and fixation output parameters are used to characterize visual performance in children with and without visual impairments from 6 months of age.

Visual problems that occur early in life can have major impact on a child's development. Without verbal communication and only based on observational ...

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer scale. However, current methods mostly rely on tethered cable recordings or only use unidirectional systems, allowing either recording or stimulation of neural activity, but not at the same time or same target. Here, a new wireless, bidirectional device for simultaneous multichannel recording and stimulation of neural activity in freely behaving rats is described. The system operates through a single portable head stage that both transmits recorded activity and can be targeted in real-time for brain stimulation using a telemetry-based multichannel software. The head stage is equipped with a preamplifier and a rechargeable battery, allowing stable long-term recordings or stimulation for up to 1 h. Importantly, the head stage is compact, weighs 12 g (including battery) and thus has minimal impact on the animal&#180;s behavioral repertoire, making the method applicable to a broad set of behavioral tasks. Moreover, the method has the major advantage that the effect of brain stimulation on neural activity and behavior can be measured simultaneously, providing a tool to assess the causal relationships between specific brain activation patterns and behavior. This feature makes the method particularly valuable for the field of deep brain stimulation, allowing precise assessment, monitoring, and adjustment of stimulation parameters during long-term behavioral experiments. The applicability of the system has been validated using the inferior colliculus as a model structure.

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

A wireless, bidirectional system for multi-channel neural recordings and stimulation in freely behaving rats is introduced. The system is light and compact, thus having minimal impact on the animal&#180;s behavioral repertoire. Moreover, this bidirectional system provides a sophisticated tool to assess causal relationships between brain activation patterns and behavior.

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer ...

Stereotactically-guided Ablation of the Rat Auditory Cortex, and Localization of the Lesion in the Brain

The rat auditory cortex (AC) is becoming popular among auditory neuroscience investigators who are interested in experience-dependence plasticity, auditory perceptual processes, and cortical control of sound processing in the subcortical auditory nuclei. To address new challenges, a procedure to accurately locate and surgically expose the auditory cortex would expedite this research effort. Stereotactic neurosurgery is routinely used in pre-clinical research in animal models to engraft a needle or electrode at a pre-defined location within the auditory cortex. In the following protocol, we use stereotactic methods in a novel way. We identify four coordinate points over the surface of the temporal bone of the rat to define a window that, once opened, accurately exposes both the primary (A1) and secondary (Dorsal and Ventral) cortices of the AC. Using this method, we then perform a surgical ablation of the AC. After such a manipulation is performed, it is necessary to assess the localization, size, and extension of the lesions made in the cortex. Thus, we also describe a method to easily locate the AC ablation postmortem using a coordinate map constructed by transferring the cytoarchitectural limits of the AC to the surface of the brain.The combination of the stereotactically-guided location and ablation of the AC with the localization of the injured area in a coordinate map postmortem facilitates the validation of information obtained from the animal, and leads to a better analysis and comprehension of the data.

We describe a method for the stereotactically-guided location, exposure, and ablation of the auditory cortex in rats. The localization of the ablation is assessed using a coordinate map postmortem.

The rat auditory cortex (AC) is becoming popular among auditory neuroscience investigators who are interested in experience-dependence plasticity, ...

A Method for Investigating Change Blindness in Pigeons (Columba Livia)

Change blindness is a phenomenon of visual attention, whereby changes to a visual display go unnoticed under certain specific circumstances. While many laboratory procedures have been developed that produce change blindness in humans, the flicker paradigm has emerged as a particularly effective method. In the flicker paradigm, two visual displays are presented in alternation with one another. If successive displays are separated by a short inter-stimulus interval (ISI), change detection is impaired. The simplicity of the procedure and the clear, performance-based operational definition of change blindness make the flicker paradigm well-suited to comparative research using nonhuman animals. Indeed, a variant has been developed that can be implemented in operant chambers to study change blindness in pigeons. Results indicate that pigeons, like humans, are worse at detecting the location of a change if two consecutive displays are separated in time by a blank ISI. Furthermore, pigeons&#39; change detection is consistent with an active, location-by-location search process that requires selective attention. The flicker task thus has the potential to contribute to investigations of the dynamics of pigeons&#39; selective spatial attention in comparison to humans. It also illustrates that the phenomenon of change blindness is not exclusive to humans&#39; visual perception, but may instead be a general consequence of selective attention. Finally, while the useful aspects of attention are widely appreciated and understood, it is also important to acknowledge that they may be accompanied by specific imperfections such as change blindness, and that these imperfections have consequences across a wide range of contexts.

A Method for Investigating Change Blindness in Pigeons Columba Livia

Change blindness is a phenomenon of visual attention, whereby changes to a visual display go unnoticed under certain specific circumstances. This protocol describes a variation on the flicker paradigm for investigating change blindness that is appropriate and effective for research with pigeons.

Change blindness is a phenomenon of visual attention, whereby changes to a visual display go unnoticed under certain specific circumstances. While many ...

A Method to Study Adaptation to Left-Right Reversed Audition

An unusual sensory space is one of the effective tools to uncover the mechanism of adaptability of humans to a novel environment. Although most of the previous studies have used special spectacles with prisms to achieve unusual spaces in the visual domain, a methodology for studying the adaptation to unusual auditory spaces has yet to be fully established. This study proposes a new protocol to set-up, validate, and use a left-right reversed stereophonic system using only wearable devices, and to study the adaptation to left-right reversed audition with the help of neuroimaging. Although individual acoustic characteristics are not yet implemented, and slight spillover of unreversed sounds is relatively uncontrollable, the constructed apparatus shows high performance in a 360&#176; sound source localization coupled with hearing characteristics with little delay. Moreover, it looks like a mobile music player and enables a participant to focus on daily life without arousing curiosity or drawing attention of other individuals. Since the effects of adaptation were successfully detected at the perceptual, behavioral, and neural levels, it is concluded that this protocol provides a promising methodology for studying adaptation to left-right reversed audition, and is an effective tool for uncovering the adaptability of humans to a novel environments in the auditory domain.

The present study proposes a protocol to investigate the adaptation to left-right reversed audition achieved only by wearable devices, using neuroimaging, which can be an effective tool for uncovering the adaptability of humans to a novel environment in the auditory domain.

An unusual sensory space is one of the effective tools to uncover the mechanism of adaptability of humans to a novel environment. Although most of the ...

A Behavioral Assay for Investigating the Role of Spatial Memory During Instinctive Defense in Mice

Evolution has selected a repertoire of defensive behaviors that are essential for survival across all animal species. These behaviors are often stereotyped actions elicited in response to innately aversive sensory stimuli, but their success requires enough flexibility for adapting to different spatial environments, which can change rapidly. Here, we describe a behavioral assay to evaluate the influence of learned spatial knowledge on defensive behaviors in mice. We have adapted the widely used Barnes maze spatial memory assay to investigate how mice navigate to a shelter during escape responses to innately aversive sensory stimuli in a novel environment, and how they adapt to acute changes in the environment. This new assay is an ethological paradigm that does not require training and exploits the natural exploration patterns and navigation strategies in mice. We propose that the set of protocols described here are a powerful means of studying goal-directed behaviors and stimulus-triggered navigation, which should be of interest to both the fields of instinctive behaviors and spatial memory.

This protocol describes a behavioral assay, based on the Barnes maze test, to study how instinctive defensive actions are modified by the knowledge of spatial environment.