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><tbody><tr><td>5-pin Connector</td><td>NW3</td><td>HD127K</td><td>1.27 mm (.050&quot;) 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>&amp;lambda; typ. 470 nm, Viewing angle 80 &amp;deg;, 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, &amp;Oslash; 200 &amp;micro;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, &amp;Oslash; 101.6 &amp;mu;m (.004&quot;)</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&amp;trade;</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>&amp;Oslash; 200 &amp;micro;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

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JoVE Journal

Neuroscience

3.7K Views.  Ewha Womans University. Dit protocol omvat het fabriceren op optrode-systeem voor gelijktijdige optogenetische stimulatie en elektrofysiologische registratie. Het voorgestelde LED-gebaseerde systeem verbetert de efficiëntie van de lichtkoppeling via een microlensarray. Een LED-lichtbron heeft een eenvoudige verlichtingsinstallatie dan een laserlichtbron, zodat het systeem gemakkelijk toepasbaar is op het draadloze systeem.Optogenetica is een padtechniek voor het begrijpen van de pathologie van neurologische ...

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

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
	One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
	Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
	We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Although the brain represents less than 5% of the body  by mass, it utilizes approximately one quarter of the glucose used by the body  at rest1. The ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

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

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

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

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

Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the identification of the neuronal type in in vivo electrophysiological experiments in broad brain regions. In optogenetics experiments, it is critical to deliver the light to the recording site. However, it is often hard to deliver the stimulation light to the deep brain regions from the brain&#39;s surface. Especially, it is difficult for the stimulation light to reach the deep brain regions when the optical transparency of the brain surface is low, as is often the case with recordings from awake animals. Here, we describe a method to record spike responses to the light from an awake mouse using a custom-made glass optrode. In this method, the light is delivered through the recording glass electrode so that it is possible to reliably stimulate the recorded neuron with light in the deep brain regions. This custom-made optrode system consists of accessible and inexpensive materials and is easy to assemble.

This work introduces a method to perform an optogenetic single-unit recording reliably from an awake mouse using a custom-made glass optrode.

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the ...

Hybrid Microdrive System with Recoverable Opto-Silicon Probe and Tetrode for Dual-Site High Density Recording in Freely Moving Mice

Multi-regional neural recordings can provide crucial information to understanding fine-timescale interactions between multiple brain regions. However, conventional microdrive designs often only allow use of one type of electrode to record from single or multiple regions, limiting the yield of single-unit or depth profile recordings. It also often limits the ability to combine electrode recordings with optogenetic tools to target pathway and/or cell type specific activity. Presented here is a hybrid microdrive array for freely moving mice to optimize yield and a description of its fabrication and reuse of the microdrive array. The current design employs nine tetrodes and one opto-silicon probe implanted in two different brain areas simultaneously in freely moving mice. The tetrodes and the opto-silicon probe are independently adjustable along the dorsoventral axis in the brain to maximize the yield of unit and oscillatory activities. This microdrive array also incorporates a set-up for light, mediating optogenetic manipulation to investigate the regional- or cell type-specific responses and functions of long-range neural circuits. In addition, the opto-silicon probe can be safely recovered and reused after each experiment. Because the microdrive array consists of 3D-printed parts, the design of microdrives can be easily modified to accommodate various settings. First described is the design of the microdrive array and how to attach the optical fiber to a silicon probe for optogenetics experiments, followed by fabrication of the tetrode bundle and implantation of the array into a mouse brain. The recording of local field potentials and unit spiking combined with optogenetic stimulation also demonstrate feasibility of the microdrive array system in freely moving mice.

This protocol describes the construction of a hybrid microdrive array that allows implantation of nine independently adjustable tetrodes and one adjustable opto-silicon probe in two brain regions in freely moving mice. Also demonstrated is a method for safely recovering and reusing the opto-silicon probe for multiple purposes.

Multi-regional neural recordings can provide crucial information to understanding fine-timescale interactions between multiple brain regions. However, ...

Automated Multimodal Stimulation and Simultaneous Neuronal Recording from Multiple Small Organisms

Fluorescent genetically encoded calcium indicators have contributed greatly to our understanding of neural dynamics from the level of individual neurons to entire brain circuits. However, neural responses may vary due to prior experience, internal states, or stochastic factors, thus generating the need for methods that can assess neural function across many individuals at once. Whereas most recording techniques examine a single animal at a time, we describe the use of wide-field microscopy to scale up neuronal recordings to dozens of Caenorhabditis elegans or other sub-millimeter-scale organisms at once. Open-source hardware and software allow great flexibility in programming fully automated experiments that control the intensity and timing of various stimulus types, including chemical, optical, mechanical, thermal, and electromagnetic stimuli. In particular, microfluidic flow devices provide precise, repeatable, and quantitative control of chemosensory stimuli with sub-second time resolution. The NeuroTracker semi-automated data analysis pipeline then extracts individual and population-wide neural responses to uncover functional changes in neural excitability and dynamics. This paper presents examples of measuring neuronal adaptation, temporal inhibition, and stimulus crosstalk. These techniques increase the precision and repeatability of stimulation, allow the exploration of population variability, and are generalizable to other dynamic fluorescent signals in small biosystems from cells and organoids to whole organisms and plants.

We present a method for the flexible chemical and multimodal stimulation and recording of simultaneous neural activity from many Caenorhabditis elegans worms. This method uses microfluidics, open-source hardware and software, and supervised automated data analysis to enable the measurement of neuronal phenomena such as adaptation, temporal inhibition, and stimulus crosstalk.

Fluorescent genetically encoded calcium indicators have contributed greatly to our understanding of neural dynamics from the level of individual neurons ...

Flexible Multi-Region Neural Recording Device for Epilepsy Research

An Integrated Method for Crafting Flexible and Convenient Electrophysiological Optrodes for Multi-Region In Vivo Recording

Epilepsy is a neurological disorder characterized by synchronized abnormal discharges involving multiple brain regions. Focal lesions facilitate the propagation of epileptic signals through associated neural circuits. Therefore, in vivo recording of local field potential (LFP) from the critical brain regions is&#160;essential for deciphering the circuits involved in seizure propagation. However, current methods for electrode fabrication and implantation lack flexibility. Here, we present a handy device designed for electrophysiological recordings (LFPs and electroencephalography [EEG]) across multiple regions. Additionally, we seamlessly integrated optogenetic manipulation and calcium signaling recording with LFP recording. Robust after-discharges were observed in several separate regions during epileptic seizures, accompanied by increasing&#160;calcium signaling. The approach used in this study offers a convenient and flexible strategy for synchronous neural recordings across diverse regions of the brain. It holds the potential for advancing research on neurological disorders by providing insights into the neural profiles of multiple regions involved in these disorders.

Author Spotlight: Unraveling Neural Communication and Circuit Interactions in Health and Disease

We have developed a simplified and cost-effective approach for electrode fabrication and conducted recordings of signals across multiple regions in freely moving mice. Utilizing optogenetics, alongside multi-region electrophysiology and calcium signal recording, enabled the revelation of neuronal activities across regions in the seizure kindling model.

Epilepsy is a neurological disorder characterized by synchronized abnormal discharges involving multiple brain regions. Focal lesions facilitate the ...

Multiscale Investigations of Cortical Processing by Integrating Laminar Polytrodes and Optogenetics with Micro Electrocorticography in Rodents

Electrocorticography (ECoG) is a methodological bridge between basic neuroscience and understanding human brain function in health and disease. ECoG records neurophysiological signals directly from the cortical surface at millisecond temporal resolution and columnar spatial resolution over large regions of cortical tissue simultaneously, making it uniquely positioned to study both local and distributed cortical computations. Here, we describe the design of custom, high-density micro-ECoG (&#181;ECoG) devices and their use in two procedures. These grids have 128 low-impedance electrodes with 200 &#181;m spacing fabricated on a clear polymer substrate with perforations between electrodes; these features enable simultaneous &#181;ECoG recording with laminar polytrode recordings and optogenetic manipulations. First, we present a protocol for combined epidural &#181;ECoG recording over the whisker somatosensory cortex of mice with optogenetic manipulation of specific genetically defined cortical cell types. This allows causal dissection of the distinct contributions of different neuronal populations to sensory processing while also monitoring their specific signatures in &#181;ECoG signals. Second, we present a protocol for acute experiments to record neural activity from the rat auditory cortex using &#181;ECoG grids and laminar polytrodes. This allows detailed topographic mapping of sensory-evoked neural responses across the cortical surface simultaneously with recordings from multiple neural units distributed across the cortical depth. These protocols enable experiments that characterize distributed cortical activity and may contribute to understanding and eventual interventions for diverse neurological disorders.

Here, we present two protocols for high-density micro-electrocorticography (&#181;EcoG) recording in rats and mice, including surgical, implantation, and recording methods. &#181;ECoG recordings are performed in combination with either laminar polytrode recording in the rat auditory cortex or with optogenetic manipulation of neural activity in the mouse somatosensory cortex.

Electrocorticography (ECoG) is a methodological bridge between basic neuroscience and understanding human brain function in health and disease. ECoG ...

Simultaneous Optical and Electrophysiological Monitoring of Neuronal Cells in Brain Slices

Source:&#160; Murphy, C. A., et al. <a href="https://app.jove.com/v/61333">Optogenetic activation of afferent pathways in brain slices and modulation of responses by volatile anesthetics.</a> J. Vis. Exp. (2020)
This video outlines a method for simultaneously imaging and recording electrical activity from transgenic brain slices that contain neuron-specific fluorescent proteins. The procedure involves maintaining the brain slice in a perfusion chamber with aerated artificial cerebrospinal fluid, using a light source to excite fluorescent-labeled cells, and recording the electrical signals with a multichannel probe and patch clamp.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.&#160;&#160;&#160;&#160;&#160;
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Optrode Array for Simultaneous Optogenetic Modulation and Electrical Neural Recording

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