This research was supported by Convergent Technology R&#38;D Program for Human Augmentation through the National Research Foundation of Korea (NRF), funded by the Ministry of Science and ICT (NRF-2019M3C1B8090805), and supported by a National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) (no. 2019R1A2C1088909). We thank Seung-Hee Lee&#39;s laboratory at the Department of Biological Sciences, KAIST, Daejeon, Korea, for kindly providing the transgenic mice.

The animal care and surgical procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at the Ewha Womans University (no. 20-029).
1. Preparation of an optrode array (Figure 1 and Figure 2)
<ol>
	<li>Attach optical fibers with the microlens array.
	<ol>
		<li>Remove the passivation coating of the optical fiber and cut it into 5 mm-long pieces using a precision optical fiber cleaver.</li>
		<li>Dip the optical fiber in the clear UV resin and place the optical fibers on the microlens array.</li>
		<li>Cure the resin using a UV lamp.</li>
		<li>Attach the microlens array to the 3D printed housing using epoxy.</li>
	</ol>
	</li>
	<li>Align perfluoroalkoxy (PFA)-coated tungsten wires.
	<ol>
		<li>Solder the 4 pieces of 30 mm-long PFA tungsten wires and 1 silver wire to the 5-pin, 1.27 mm pitch female connector. Use silver wire as a reference electrode.</li>
		<li>Connect the 1.27 mm pitch female connector to the headstage preamplifier.</li>
		<li>Carefully put tungsten wires through the silicon via the holes (Figure 1B) to help the fine alignment of the tungsten electrodes.</li>
		<li>Cut the aligned tungsten wires 1 mm shorter than the optical fiber length.</li>
		<li>Attach the connector to the 3D printed housing using epoxy.</li>
	</ol>
	</li>
	<li>LED placement
	<ol>
		<li>Place the LED in the 3D printed housing.</li>
		<li>Connect the LED to the driving circuit that generates pulse width modulation (PWM). 
		NOTE: The PWM pulse is generated by the microcontroller.</li>
		<li>Given the noise induced by the LED driving circuit, ensure the recording electrodes are 50 mm apart from the LED driving circuit.</li>
		<li>Measure the light intensity at the end of the optical fiber tip using a photodiode.</li>
	</ol>
	</li>
	<li>Immerse the tungsten electrodes and optical fibers in alcohol for 15 min.&#160;Then immerse the system in sterile D.I water and sterilize with ethylene oxide gas.</li>
</ol>
2. Implantation surgery (Figure 3 and Figure 4)
NOTE: Sterile technique must be followed during&#160;surgery.
<ol>
	<li>Prepare the transgenic animals, which are genetically modified to express light-sensitive opsin proteins in the specific type of neuronal cells. 
	NOTE: A male, 2 months old, transgenic mouse expressing channelrhodopsin-2 (ChR2) in Ca2+/calmodulin-dependent protein kinase II-positive neurons (CaMKII&#945;::ChR2 mouse) was used in this study41.</li>
	<li>Anesthetize the mouse with a ketamine-xylazine cocktail-a mixture of ketamine (100 mg/kg) and xylazine (10 mg/kg)-by intraperitoneal administration.
	<ol>
		<li>Check the anesthetized animal every 30 min by monitoring whisker movement and reaction to paw pinch.</li>
		<li>Inject the ketamine-xylazine cocktail at half of the initial dose if additional anesthesia is needed.</li>
	</ol>
	</li>
	<li>Shave the head skin and place the anesthetized mouse in the stereotactic frame. 
	NOTE: Perform surgical skin prep and drape animal. Skin preparation involves hair removal followed by cleaning of the&#160;skin with antimicrobial agents.
	<ol>
		<li>Position the head within the stereotactic frame and insert ear bars into the meatus.</li>
		<li>Center the mouse head in the stereotactic frame by repeatedly loosening and tightening the ear bars for exact positioning.</li>
		<li>Place the incisor bar so that the upper incisors hook over the front inside edge.</li>
		<li>Adjust the incisor bar for setting the height of the incisor bar, and tighten the nose clamp against the snout.</li>
		<li>Turn on the thermal heating pad embedded in the stereotactic frame in advance to maintain the body temperature at 37 &#176;C throughout the surgical procedures.</li>
		<li>Put the thermal probe into the mouse&#39;s rectum for homeothermic regulation. 
		NOTE: Ear bar placement must be checked by gently grasping and swaying the snout. The ear bar must be readjusted if the snout can be moved more than ~4 mm laterally.</li>
	</ol>
	</li>
	<li>Cover the eyes with petroleum jelly (see the Table of Materials) to prevent dryness.</li>
	<li>Inject 1% lidocaine into the scalp. Lift the skin of the head with forceps, secure an injection space, and inject the lidocaine under the scalp.</li>
	<li>Perform a sagittal incision using a scalpel and fine scissors. Hold the incised skin with a microclamp to broaden the visibility of the surgical area. 
	NOTE: The incision length is &#60;1 cm above the bregma and caudal edge of the interparietal bone.</li>
	<li>Remove the periosteum using cotton swabs. If there is bleeding, cauterize the skull using a bovie to seal the blood vessels.</li>
	<li>Clean the skull with saline and mark the craniotomy sites using a manipulator arm: hippocampus Anterior-Posterior (AP) &#8722;1.8 to &#8722;2.8 mm, Medial-Lateral (ML) 0.5-2.5 mm, and Dorsal-Ventral (DV) &#8722;1 to &#8722;2 mm. 
	NOTE: Considering the thickness of the mouse skull, insert the optrode array -1.2 to -2.2 mm from the exposed brain area. Given the hippocampal pathway, pyramidal cells of CA3 send their axons to CA1. Therefore, optical fibers are 1 mm longer than recording electrodes, so that the optical fibers are placed in CA3 and the recording electrodes in CA1.</li>
	<li>Drill a hole above the cerebellum and insert a screw for ground. Put the ground screw 0.5 mm deep with a precision screw until it reaches the top of the cerebellum. 
	NOTE: Ensure tight insertion of the screw, which will be used as a ground electrode and a supporting structure to maximize the long-term stability of implantation, as it increases the three-dimensional surface for adherence of the dental cement.</li>
	<li>Drill the marked area and remove a piece of the skull with forceps.</li>
	<li>Bend a 26 G needle tip to clockwise 120&#176;, and expose the brain area by inserting the bevel side of the needle faced upward. Be careful not to damage the brain and blood vessels.</li>
	<li>Clean the exposed brain with saline to wash out bone dust and extraneous matter.</li>
	<li>Fix the optrode array in the manipulator arm and move close to the exposed area. 
	NOTE: When placing the device, recalculating the target area again from the bregma can increase the accuracy of the placement.</li>
	<li>Slowly insert the optrode array and connect the ground screw to the silver wire attached to the system42. 
	NOTE: In this process, the array must be tightly fixed to the manipulator arm to minimize wobbling to prevent brain damage from micromotion during insertion. Insertion must be performed slowly, with rest for 10 min to account for swelling of the brain that may have been pressed after insertion. Insertion speed under 1 &#181;m/s into the brain tissue is recommended for the high quality of the signal-to-noise ratio and the number of separable single units42.</li>
	<li>If there is bleeding, apply direct pressure to bleeders with a dry cotton swab or use&#160;cautery. Do not move on to the next steps until bleeding is completely stopped.</li>
	<li>Insert gel foam between the exposed brain and the device to protect chemicals from direct contact with the brain. 
	NOTE: Gel foam also helps hemostasis and keeps the brain moist.</li>
	<li>Wipe the skull with cotton swabs to remove moisture as much as possible, and proceed to the next fixation step.</li>
	<li>Carefully apply dental cement to the skull to fix the device and cover the gel foam. Before the dental cement is completely hardened, separate the scalp if attached to dental cement. Pull the incised skin with forceps to cover the hardened dental cement and suture the scalp. Then, release the device from the manipulator arm. 
	&#8203;NOTE: Ensure that the dental cement does not stick to the skin and only covers the skull region. When the skin grows, it can push the cement away and, eventually, the device will fall off. This will cause a critical problem for long-term in vivo experiments.</li>
</ol>
3. Recovery and implant care
<ol>
	<li>Take the mouse out of the stereotactic frame.</li>
	<li>Administer Carprofen solution using a 26 G needle. Inject once before surgery and 12 h (next morning in case of afternoon surgery) and 24 h after surgery to reduce pain. 
	NOTE: The drug concentration in the bottle is 50 mg/mL, and it is stored at 4 &#176;C. Carprofen is viscous and needs to be diluted in sterile water 1:10. If sterility is maintained, the solution can be stored for up to 4 weeks. To maintain the effective duration of the drug, inject the Carprofen solution at 12 h intervals.</li>
	<li>Place the mouse on the heating pad and check if it recovers well from the anesthesia. 
	NOTE: The animal is not left unattended until it regains sufficient consciousness.</li>
	<li>Remove the suture threads 7 - 10 days after surgery.</li>
	<li>Allow the animal to recover for 1 week. Monitor food and water intake during recovery time. Administer the analgesic and check for the signs of discomfort or pain. 
	NOTE: The animals that have undergone surgery cannot stay with the other animals until they fully recover.
	<ol>
		<li>During the recovery period, weigh the mouse every day to check for weight changes. Sacrifice the mouse (see section 6) if there is 20% weight loss compared to age and sex matched control animals.</li>
	</ol>
	</li>
</ol>
4. Optogenetic stimulation and electrophysiological recording
<ol>
	<li>Anesthetize the mouse and place the anesthetized mouse in the stereotactic frame.</li>
	<li>Set up the stimulation parameters.
	<ol>
		<li>Set the light pulse recipe to 4% duty cycle and 10 Hz frequency43. Set up the light stimulation for 2 s (20 pulses) during neural recording.</li>
		<li>Use a 50 mA current to drive 3 mW/mm2 light intensity at the optical fiber tip. Measure the light power using a photodiode and power meter and divide the light power by the optical fiber facet area.</li>
	</ol>
	</li>
	<li>Take off the plastic cover and attach the upper part containing the light delivery system with the reusable LED and circuits.</li>
	<li>Connect the headstage preamplifier to the implanted 5 pin connector.</li>
	<li>Open the software to record neural signals.</li>
	<li>Set up the software filters. Use a bandpass filter at 0.1-20 kHz and a notch filter at 60 Hz. Set the amplifier sampling rate to 20 kHz.</li>
	<li>Before the recording, check if the neural spike signals are detected. 
	NOTE: Noise cancellation is important because neural signals are too small. If the noise level is too high, neural signals may be obscured by noise and become invisible. Use appropriate grounding and electromagnetic wave shielding to reduce noise.</li>
	<li>After the signal check, record neural signals without light stimulation for baseline recording.</li>
	<li>Deliver LED light and simultaneously record neuronal responses (Figure 5 and Figure 6). 
	&#8203;NOTE: Once stimulated, the next stimulation experiment must be performed after an interval of at least 5 min so that the neuronal activity evoked by the previous stimulation does not affect the next experiment.</li>
</ol>
5. Data analysis
<ol>
	<li>Load the acquired data using MATLAB program.</li>
	<li>Obtain single neural activities using a spike sorting algorithm &#34;Wave-Clus&#34;44,45. Detect the spikes in each channel by an amplitude threshold as in Eq (1)45. 
	Threshold = 5 &#215; median <img alt="Equation 1" src="/files/ftp_upload/63460/63460eq01.jpg" /> (1) 
	&#8203;Where x is the bandpass-filtered signal.
	<ol>
		<li>To install &#34;Wave-Clus,&#34; refer to the download link in the Table of Materials.</li>
		<li>To open the graphical user interface (GUI), type wave_clus in the command prompt of MATLAB.</li>
		<li>Type Get_spikes(&#39;filename.ext&#39;) function for file extension prepared to spike detection. Then, run Do_clustering(&#39;filename_spikes.mat&#39;) for spike sorting.</li>
	</ol>
	</li>
	<li>Count the sorted spikes before, during, and after light stimulation with specific time bins. Use 0.2 s and 2 s time bins for analysis.</li>
	<li>Plot the number of spikes from each recording channel. 
	&#8203;NOTE: Mean with standard error of the means (SEM) of the data from the results of 8 repeated experiments was calculated and plotted.</li>
</ol>
6. Euthanasia
<ol>
	<li>After all the experiments, sacrifice the mouse by carbon dioxide (CO2) inhalation.</li>
	<li>Without precharging the chamber, place the mouse in the transparent euthanasia chamber, without CO2 supply and/or leaks.</li>
	<li>Turn on the CO2 gas, and displace the air at a rate of 30 - 50% of the volume of euthanasia chamber air per min. 
	NOTE: A flowmeter must be connected to the CO2 gas cylinder to ensure that the air displacement rate is 30-50% of the volume of the euthanasia chamber per min.</li>
	<li>Monitor the sacrificed mouse for lack of respiration and changed eye color. 
	NOTE: The expected time for euthanasia is usually within 10 to 15 min.</li>
	<li>After observing the death signs, remove the mouse from the euthanasia chamber.</li>
	<li>To complete the euthanasia procedure, verify death by confirming respiratory and cardiac arrest.</li>
</ol>

The authors have no conflicts of interest to disclose.

The feasibility of the system for simultaneous optogenetic stimulation and electrophysiological recording was verified (Figure 6). The big spikes during light stimulation are photoelectric artifacts occurring at the same time as the light stimulation (Figure 6A). This is clear in the zoomed view of the waveform in the red dashed rectangle (Figure 6A). As shown in Figure 6A, the photoelectrical artifacts...

Recording and controlling neural activity is essential for understanding how the brain functions in a neural network and at cellular levels. Conventional electrophysiological recording methods include the patch clamp1,2,3,4 using a micropipette and extracellular recording using microneural electrodes5,6,7,8. As a neuromodulation method, electrical stimulation has been frequently used to directly stimulate a focal brain region through direct or indirect depolarization of neuronal cells. However, the electrical method cannot distinguish neuronal cell types for recording or stimulation because the electrical currents spread in all directions.
As an emerging technology, optogenetics has ushered in a new era in understanding how the nervous system works9,10,11,12,13,14,15,16. The essence of optogenetic techniques is to use light to control the activity of light-sensitive opsin proteins expressed by genetically modified cells. Thus, optogenetics enables the sophisticated modulation or monitoring of genetically selected cells in complicated neural circuits14,17. The wider use of the optogenetic approach has necessitated simultaneous neural recording to directly confirm optical neuromodulation. Therefore, an integrated device with light control and recording functions would be extremely valuable16,18,19,20,21,22,23,24,25.
There are limitations of conventional, laser-based optogenetic stimulation, which requires a bulky and expensive light delivery system26,27,28,29,30. Therefore, some research groups employed &#181;LED-based silicon probes to minimize the size of the light delivery system31,32,33,34. However, there is a risk of thermal brain damage caused by direct contact with &#181;LEDs due to the low energy conversion efficiency of LEDs. Light waveguides, such as optical fibers, SU-8, and silicon oxynitride (SiON), have been applied to avoid thermal damage30,35,36,37,38,39. However, this strategy also has a drawback due to its low coupling efficiency between light sources and the waveguides.
The microlens array was previously introduced to enhance the light coupling efficiency between LEDs and optical fibers40. An optrode system was developed based on microelectromechanical systems (MEMS) technologies for optical stimulation and electrical recording on a microscale40. The microlens array between an LED and optical fibers increased the light efficiency by 3.13 dB. As shown in Figure 1, a 2x2 optical fiber array is aligned on the 4x4 microlens array, and the LED is positioned below the microlens array. The 2x2 optical fibers are mounted instead of 4x4 to reduce brain damage. A tungsten electrode array is positioned adjacent to the optrode array using silicon via holes for electrophysiological recording (Figure 1B).
The system consists of a top disposable part and detachable bottom parts. The top disposable part, which includes the optical fiber array, microlens array, and the tungsten electrode array, is designed to be permanently implanted into the brain for in vivo experiments. The bottom part includes an LED light source and an external power supply line, which is easily removable and reusable for another animal experiment. An attachable plastic cover protects the disposable part when the detachable part is removed.
The feasibility of the system is verified by implantation into the brains of transgenic mice expressing channelrhodopsin-2 (ChR2) in Ca2+/calmodulin-dependent protein kinase II-positive neurons (CaMKII&#945;::ChR2 mouse). Recording electrodes were used to record the neural activities from individual neurons during optical stimulation of the neurons.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>5-pin Connector</td>
			<td>NW3</td>
			<td>HD127K</td>
			<td>1.27 mm (.050&#34;) pitch</td>
		</tr>
		<tr>
			<td>Bovie</td>
			<td>Fine Science Tools(F.S.T)</td>
			<td>18010-00</td>
			<td>High Temperature Cautery Kit</td>
		</tr>
		<tr>
			<td>Data Acquisition Software</td>
			<td>Intan Technologies, LLC</td>
			<td>USB Interface Board software</td>
			<td>Work with the RHD USB Interface Board</td>
		</tr>
		<tr>
			<td>Dental Cement</td>
			<td>Lang Dental Manufacturing Company, Inc.</td>
			<td>1223CLR</td>
			<td>Use Jet Liquid and powder in jet denture repair package</td>
		</tr>
		<tr>
			<td>Digital Manipulator Arm</td>
			<td>Stoelting Co.</td>
			<td>51904/51906</td>
			<td>Left, Right each Digital Manipulator Arm, 3-Axes, Add-On</td>
		</tr>
		<tr>
			<td>Gel Foam</td>
			<td>Cutanplast</td>
			<td>Standard (70*50*10 mm)</td>
			<td>Sterile re-absorbable gelatin sponge with a haemostatic effect</td>
		</tr>
		<tr>
			<td>Headstage Preamplifier</td>
			<td>Intan Technologies, LLC</td>
			<td>#C3314</td>
			<td>RHD 16-Channel Recording Headstages</td>
		</tr>
		<tr>
			<td>Heating Pad</td>
			<td>Stoelting Co.</td>
			<td>53800R</td>
			<td>Stoelting Rodent Warmer X1 with Rat Heating Pad</td>
		</tr>
		<tr>
			<td>LED</td>
			<td>OSLON</td>
			<td>GB CS8PM1.13</td>
			<td>&#955; typ. 470 nm, Viewing angle 80 &#176;, Forward voltage 2.85 V</td>
		</tr>
		<tr>
			<td>MATLAB</td>
			<td>MathWorks, Inc.</td>
			<td>R2019a</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Clamp</td>
			<td>SURGIWAY</td>
			<td>12-1002-04</td>
			<td>Straight type, Serre-fine DIEFFENBACH droite 3.5 cm</td>
		</tr>
		<tr>
			<td>Optical Fiber</td>
			<td>Thorlabs, Inc.</td>
			<td>FT200UMT</td>
			<td>0.39 NA, &#216; 200 &#181;m Core Multimode Optical Fiber, High OH for 300 - 1200 nm</td>
		</tr>
		<tr>
			<td>PFA-Coated Tungsten Wire</td>
			<td>A-M System</td>
			<td>Custom ordered</td>
			<td>Rod type, &#216; 101.6 &#956;m (.004&#34;)</td>
		</tr>
		<tr>
			<td>Photodiode</td>
			<td>Thorlabs</td>
			<td>S121C</td>
			<td></td>
		</tr>
		<tr>
			<td>power meter</td>
			<td>Thorlabs Inc.</td>
			<td>PM100D</td>
			<td></td>
		</tr>
		<tr>
			<td>Precision cleaver</td>
			<td>FITEL</td>
			<td>S326</td>
			<td>Fiber slicer tool</td>
		</tr>
		<tr>
			<td>Prism</td>
			<td>GraphPad</td>
			<td>5.01 version</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Feather&#8482;</td>
			<td>#20</td>
			<td>Scalpel blade with 100mm long Scalpel Handle</td>
		</tr>
		<tr>
			<td>screw</td>
			<td>Nasa Korea</td>
			<td>stainless steel</td>
			<td>diameter: 1.2 mm, length: 3 mm</td>
		</tr>
		<tr>
			<td>Silver Wire</td>
			<td>The Nilaco Corporation</td>
			<td>AG-401265</td>
			<td>&#216; 200 &#181;m</td>
		</tr>
		<tr>
			<td>Stereotaxic Fxrame</td>
			<td>Stoelting Co.</td>
			<td>51500D</td>
			<td>Digital new standard stereotaxic, rat and mouse</td>
		</tr>
		<tr>
			<td>suture</td>
			<td>ETHICON</td>
			<td>W9106</td>
			<td>suture size: 4-0, length:75 cm, wire diameter: 4-0</td>
		</tr>
		<tr>
			<td>Vaseline</td>
			<td>Unilever PLC</td>
			<td>Original</td>
			<td>100% pure petroleum jelly</td>
		</tr>
		<tr>
			<td>Wave_Clus</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>https://github.com/csn-le/wave_clus</td>
		</tr>
	</tbody>
</table>

optrode array for simultaneous optogenetic modulation and electrical neural recording

During the last decade, optogenetics has become an essential tool for the investigation of neural signaling due to its unique capability of selective neural modulation or monitoring. As specific types of neuronal cells can be genetically modified to express opsin proteins, optogenetics enables optical stimulation or inhibition of the selected neurons. There have been several technological advances in the optical system for optogenetics. Recently, it was proposed to combine the optical waveguide for light delivery with electrophysiological recording to simultaneously monitor the neural responses to optogenetic stimulation or inhibition. In this study, an implantable optrode array (2x2 optical fibers) was developed with embedded multichannel electrodes. 
A light-emitting diode (LED) was employed as a light source, and a microfabricated microlens array was integrated to provide sufficient light power at the tip of the optical fibers. The optrode array system comprises the disposable part and the reusable part. The disposable part has optical fibers and electrodes, while the reusable part has the LED and electronic circuitry for light control and neural signal processing. The novel design of the implantable optrode array system is introduced in the accompanying video in addition to the procedure of the optrode implantation surgery, optogenetic light stimulation, and the electrophysiological neural recording. The results of in vivo experiments successfully showed time-locked neural spikes evoked by the light stimuli from hippocampal excitatory neurons of mice.

The optrode system is successfully fabricated to provide sufficient light power to activate the target neurons. The fine alignment of the tungsten electrodes is achieved through the microfabricated silicon via the holes. The measured light intensity is 3.6 mW/mm2 at the optical fiber tip when 50 mA current is applied. The microlens increased the light efficiency by 3.13 dB. Due to the microlens array, which enhances the light coupling, the applied current is approximately half of the current required to achiev...

Watch this Scientific Journal Video about Optrode Array for Simultaneous Optogenetic Modulation and Electrical Neural Recording at JoVE.com

Optrode Array for Simultaneous Optogenetic Modulation and Electrical Neural Recording

Here, we present the fabrication method of an optrode system with optical fibers for light delivery and an electrode array for neural recording. In vivo experiments with transgenic mice expressing channelrhodopsin-2 show the feasibility of the system for simultaneous optogenetic stimulation and electrophysiological recording.

During the last decade, optogenetics has become an essential tool for the investigation of neural signaling due to its unique capability of selective ...

optrode-array-for-simultaneous-optogenetic-modulation-electrical

Research

JoVE Journal

Neuroscience

3.6K Views.  Ewha Womans University. Here, we present the fabrication method of an optrode system with optical fibers for light delivery and an electrode array for neural recording. In vivo experiments with transgenic mice expressing channelrhodopsin-2 show the feasibility of the system for simultaneous optogenetic stimulation and electrophysiological recording. 

Video: Optrode Array for Simultaneous Optogenetic Modulation and Electrical Neural Recording

Design and Construction of a Cost Effective Headstage for Simultaneous Neural Stimulation and Recording in the Water Maze

Headstage preamplifiers and source followers are commonly used to study neural activity in behavioral neurophysiology experiments. Available commercial products are often expensive, not easily customized, and not submersible. Here we describe a method to design and build a customized, integrated circuit headstage for simultaneous 4-channel neural recording and 2-channel simulation in awake, behaving animals. The headstage is designed using a free, commercially available CAD-type design package, and can be modified easily to accommodate different scales (e.g. to add channels). A customized printed circuit board is built using surface mount resistors, capacitors and operational amplifiers to construct the unity gain source follower circuit. The headstage is made water-proof with a combination of epoxy, parafilm and a synthetic rubber putty. We have successfully used this device to record local field potentials and stimulate different brain regions simultaneously via independent channels in rats swimming in a water maze. The total cost is &lt; $30/unit and can be manufactured readily.

We present a low-cost method to design and construct a light headstage pre-amplifier system with simultaneous neural recording and stimulation capability. This device can be waterproofed for use in swimming animals.

Headstage preamplifiers and source followers are commonly used to study neural activity in behavioral neurophysiology experiments. Available commercial ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Optogenetic Functional MRI

The investigation of the functional connectivity of precise neural circuits across the entire intact brain can be achieved through optogenetic functional magnetic resonance imaging (ofMRI), which is a novel technique that combines the relatively high spatial resolution of high-field fMRI with the precision of optogenetic stimulation. Fiber optics that enable delivery of specific wavelengths of light deep into the brain in vivo are implanted into regions of interest in order to specifically stimulate targeted cell types that have been genetically induced to express light-sensitive trans-membrane conductance channels, called opsins. fMRI is used to provide a non-invasive method of determining the brain's global dynamic response to optogenetic stimulation of specific neural circuits through measurement of the blood-oxygen-level-dependent (BOLD) signal, which provides an indirect measurement of neuronal activity. This protocol describes the construction of fiber optic implants, the implantation surgeries, the imaging with photostimulation and the data analysis required to successfully perform ofMRI. In summary, the precise stimulation and whole-brain monitoring ability of ofMRI are crucial factors in making ofMRI a powerful tool for the study of the connectomics of the brain in both healthy and diseased states.

This protocol describes the steps and data analysis required to successfully perform optogenetic functional magnetic resonance imaging (ofMRI). ofMRI is a novel technique that combines high-field fMRI readout with optogenetic stimulation, allowing for cell type-specific mapping of functional neural circuits and their dynamics across the whole living brain.

The investigation of the functional connectivity of precise neural circuits across the entire intact brain can be achieved through optogenetic functional ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a closed-loop event-triggered manner. Although many studies are focusing on the possible benefits and side-effects of TES in healthy and pathologic brains, there are still many fundamental open questions regarding the mechanism of action of the stimulation. Therefore, there is a clear need for a robust and reproducible method to test the acute and the chronic effects of TES in rodents. TES can be combined with regular behavioral, electrophysiological, and imaging techniques to investigate neuronal networks in vivo. The implantation of transcranial stimulation electrodes does not impose extra constraints on the experimental design while it offers a versatile, flexible tool to manipulate brain activity. Here we provide a detailed, step-by-step protocol to fabricate and implant transcranial stimulation electrodes to influence brain activity in a temporally constrained manner for months.

This detailed protocol describes transcranial stimulation electrode placement on the temporal bone in order to investigate the short- and long-term effects of transcranial electrical stimulation in freely moving rats.

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a ...

Simultaneous Recordings of Cortical Local Field Potentials, Electrocardiogram, Electromyogram, and Breathing Rhythm from a Freely Moving Rat

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls body functions and internal organ rhythms when animals are exposed to emotional challenges and changes in their living environments. In general experiments, signals from different organs, such as the brain and the heart, are recorded by independent recording systems that require multiple recording devices and different procedures for processing the data files. This study describes a new method that can simultaneously monitor electrical biosignals, including tens of local field potentials in multiple brain regions, electrocardiograms that represent the cardiac rhythm, electromyograms that represent awake/sleep-related muscle contraction, and breathing signals, in a freely moving rat. The recording configuration of this method is based on a conventional micro-drive array for cortical local field potential recordings in which tens of electrodes are accommodated, and the signals obtained from these electrodes are integrated into a single electrical board mounted on the animal's head. Here, this recording system was improved so that signals from the peripheral organs are also transferred to an electrical interface board. In a single surgery, electrodes are first separately implanted into the appropriate body parts and the target brain areas. The open ends of all of these electrodes are then soldered to individual channels of the electrical board above the animal's head so that all of the signals can be integrated into the single electrical board. Connecting this board to a recording device allows for the collection of all of the signals into a single device, which reduces experimental costs and simplifies data processing, because all data can be handled in the same data file. This technique will aid the understanding of the neurophysiological correlates of the associations between central and peripheral organs.

This study introduces a method for the simultaneous recording of local field potentials in the brain, electrocardiograms, electromyograms, and breathing signals of a freely moving rat. This technique, which reduces experimental costs and simplifies data analysis, will contribute to the understanding of the interactions between the brain and peripheral organs.

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls ...

Optogenetic Manipulation of Neural Circuits During Monitoring Sleep/wakefulness States in Mice

In recent years, optogenetics has been widely used in many fields of neuroscientific research. In many cases, an opsin, such as channel rhodopsin 2 (ChR2), is expressed by a virus vector in a particular type of neuronal cells in various Cre-driver mice. Activation of these opsins is triggered by application of light pulses which are delivered by laser or LED through optic cables, and the effect of activation is observed with very high time resolution. Experimenters are able to acutely stimulate neurons while monitoring behavior or another physiological outcome in mice. Optogenetics can enable useful strategies to evaluate function of neuronal circuits in the regulation of sleep/wakefulness states in mice. Here we describe a technique for examining the effect of optogenetic manipulation of neurons with a specific chemical identity during electroencephalogram (EEG) and electromyogram (EMG) monitoring to evaluate the sleep stage of mice. As an example, we describe manipulation of GABAergic neurons in the bed nucleus of the stria terminalis (BNST). Acute optogenetic excitation of these neurons triggers a rapid transition to wakefulness when applied during NREM sleep. Optogenetic manipulation along with EEG/EMG recording can be applied to decipher the neuronal circuits that regulate sleep/wakefulness states.

Here, we describe methods of optogenetic manipulation of particular types of neurons during monitoring of sleep/wakefulness states in mice, presenting our recent work on the bed nucleus of the stria terminalis as an example.

In recent years, optogenetics has been widely used in many fields of neuroscientific research. In many cases, an opsin, such as channel rhodopsin 2 ...

Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent development of thin, flexible ECoG electrodes has enabled conduction of stable recordings of large-scale cortical activity. We have developed a whole-cortical ECoG array for the common marmoset. The array continuously covers almost the entire lateral surface of cortical hemisphere, from the occipital pole to the temporal and frontal poles, and it captures whole-cortical neural activity in one shot. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain. Marmosets have two advantages regarding ECoG recordings, one being the homologous organization of anatomical structures in humans and macaques, including frontal, parietal, and temporal complexes. The other advantage is that the marmoset brain is lissencephalic and contains a large number of complexes, which are more difficult to access in macaques with ECoG, that are exposed to the brain surface.These features allow direct access to most cortical areas beneath the surface of the brain. This system provides an opportunity to investigate global cortical information processing with high resolutions at a sub-millisecond order in time and millimeter order in space.

We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Optogenetic Activation of Afferent Pathways in Brain Slices and Modulation of Responses by Volatile Anesthetics

Anesthetics influence consciousness in part via their actions on thalamocortical circuits. However, the extent to which volatile anesthetics affect distinct cellular and network components of these circuits remains unclear. Ex vivo brain slices provide a means by which investigators may probe discrete components of complex networks and disentangle potential mechanisms underlying the effects of volatile anesthetics on evoked responses. To isolate potential cell type- and pathway-specific drug effects in brain slices, investigators must be able to independently activate afferent fiber pathways, identify non-overlapping populations of cells, and apply volatile anesthetics to the tissue in aqueous solution. In this protocol, methods to measure optogenetically-evoked responses to two independent afferent pathways to neocortex in ex vivo brain slices are described. Extracellular responses are recorded to assay network activity and targeted whole-cell patch clamp recordings are conducted in somatostatin- and parvalbumin-positive interneurons. Delivery of physiologically relevant concentrations of isoflurane via artificial cerebral spinal fluid to modulate cellular and network responses is described.

Ex vivo brain slices can be used to study the effects of volatile anesthetics on evoked responses to afferent inputs. Optogenetics are employed to independently activate thalamocortical and corticocortical afferents to non-primary neocortex, and synaptic and network responses are modulated with isoflurane.

Anesthetics influence consciousness in part via their actions on thalamocortical circuits. However, the extent to which volatile anesthetics affect ...