The authors were supported by the Japan Society for the Promotion of Science KAKENHI Grant JP16K11200 and 17H02223, and the Grant for Research from Kanazawa Medical University S2016-8 and C2017-3. We thank Yuhichi Kuda for his support in taking the photos.

All procedures are done in accordance with the guiding principles of the Physiological Society of Japan and with the approval of the Animal Care Committee of Kanazawa Medical University.
1. Construction of the Glass Optrode Holder
NOTE: To build a glass optrode holder, a commercial electrode holder is used (Figure 1A).
<ol>
	<li>Gently pull out the steel tube for pressure control from the barrel of the holder.</li>
	<li>By drilling, broaden the hole of the pin seat side in the barrel of the holder. Make the hole 3 mm in diameter and 12 mm in depth.</li>
	<li>Put a stainless-steel pipe (3 mm in diameter, 0.5 mm in thickness, and 8 mm in length; Figure 1A2) in the hole.</li>
	<li>Insert a ceramic split mating sleeve for &#8960;2.5 mm ferrules (Figure 1A1) into the hole.</li>
	<li>Fix the barrel of the electrode holder to an L-shaped rabbet with epoxy adhesive.</li>
	<li>Make the holes (3 mm in diameter) in the rabbet and attach it to a manipulator with screws.</li>
</ol>
2. Head Post Installation
<ol>
	<li>Prepare a custom-built head post made of a circuit board spacer. Make the columnar circuit board spacer into a cuboidal shape with a milling cutter and cut it into 2 head posts with a saw (Figure 1B). Flatten the bottom of the head post with a milling cutter.</li>
	<li>Prepare the transgenic animal in which a light-sensitive opsin is expressed in a specific neuron type for the study. In this work, the transgenic mouse (VGAT-ChR2 mice) in which the inhibitory neurons express channelrhodopsin-2 (ChR2)11 is used.</li>
	<li>Anesthetize the animal with a mixture of 0.3 mg/kg of medetomidine, 4.0 mg/kg of midazolam, and 5.0 mg/kg of butorphanol by intraperitoneal administration. To confirm that the animal is fully anesthetized, test its reactions to tail pinching.</li>
	<li>Place the mouse in a stereotaxic frame on a heating pad. Clean the stereotaxic frame and the heating pad with 70% alcohol in advance. Place the teeth in the hole of the bite bar, and lightly tighten the nose clamp. Fix both sides of the head using ear bars. When the head is correctly fixed by the ear bars, tighten the nose clamp. Apply an ophthalmic ointment to the eyes.</li>
	<li>Shave the head skin with small scissors and clean the scalp with a cotton swab of chlorhexidine gluconate. Before the surgery, sterilize all the surgical instruments and a head post with an autoclave (121 &#176;C, 15 min).</li>
	<li>Cut the scalp along the midline with scissors and push it aside. Make the incision from the back of the head to the eyes (approximately 20 mm). Remove the periosteum with a cotton swab dipped in 70% alcohol and dry the skull.</li>
	<li>Attach the head post to the manipulator. Position the head post flat on the skull around the bregma using the manipulator.</li>
	<li>Mix the monomer (4 drops), catalyst (1 drop), and polymer (1 small spoon) of dental cement on a small dish that is chilled in the refrigerator in advance. Put the dental cement (below 1 g) on the frontal and parietal bone of the skull to fix the head post to the skull.</li>
	<li>After the dental cement becomes hard, remove the mouse from the stereotaxic frame. Put an antibiotic ointment on the exposed skin. Inject the medetomidine antagonist (0.3 mg/kg) and the antibiotics (oxytetracycline; 20 mg/kg). Place and monitor the mouse on the heat pad in a cage until it awakes, and return it to a housing cage.</li>
	<li>House the animal in an individual housing cage with some enrichment device. Change the animal paper bedding daily to keep the cage clean and prevent infection. Carefully monitor whether the animal shows any sign of discomfort. Apply lidocaine to the wound if any pain is suspected.&#160;Administer meloxicam or carprofen once every 24 h for 48 h after both the head post installation and craniotomy surgeries. The local lidocaine can be given in addition if the NSAID is not adequate.</li>
</ol>
3. Acclimation
<ol>
	<li>Two days (minimum) after the recovery from the anesthesia, acclimate the animals to the head-fixed state in the recording chamber. Perform the acclimation sessions for at least 5 days.</li>
	<li>During the acclimation sessions, place the animal in the recording chamber with the head post fixed to a custom-built clamp. During the session, in order to limit movement, cover the body with a custom-made resin half tube (Figure 1C), made using a 3D printer. 
	NOTE: On day 1, the acclimation session should be 5 min long. It is then increased daily to 10, 20, 40, and 60 min for days 2, 3, 4, and 5, respectively.</li>
	<li>At the end of each session, give the animal a food reward. If the animal struggles continuously during the acclimation, stop the session.</li>
</ol>
4. Craniotomy
NOTE: After the acclimation, a craniotomy is made over the brain region for recording. The craniotomy is performed in the stereotaxic frame under anesthesia, as with the head post installation. The post-operation procedure is the same as the head post installation.&#160;
<ol>
	<li>Before the craniotomy, clean the skull with a cotton swab with chlorhexidine gluconate and 70% alcohol.</li>
	<li>Carefully scrape the skull over the recording site with a dental drill. When the bone becomes thin enough, cut it with the tip of the scalpel and remove it.</li>
	<li>Put dental cement around the exposed region in order to strengthen the skull.</li>
	<li>Cover the exposed brain area with silicone adhesive after the dental cement becomes hard. The post-surgery procedure is as described in step 2.8.</li>
</ol>
5. Electrophysiological Recording
<ol>
	<li>Allow the animal to recover for at least 1 day after the craniotomy before performing the electrophysiological recording. Place the animal in the recording chamber in the same way it was done during the acclimation sessions when the recording starts.</li>
	<li>Make glass pipettes from borosilicate glass capillaries (OD = 1.5 mm, ID = 0.9 mm, 90.0 mm long) pulled on an electrode puller. Their tip diameter is 2&#8211;3 &#181;m, and their resistance is 4&#8211;6 M&#937; when they are filled with 10 mM PBS. The length of the electrode is 40&#8211;50 mm. In some recordings, 2% neurobiotin is added to the 10 mM PBS.</li>
	<li>Attach the filled glass pipette to the custom-made optrode holder.</li>
	<li>Put the Ag-Cl coated silver wire (0.2 mm in diameter) into the glass pipette. Connect the silver wire to a pin via a thin coated electric wire. Connect the pin to the headstage of the recording amplifier.</li>
	<li>Connect the fiber-optic patch cord (core = 960 &#181;m, cladding = 1,000 &#181;m, NA = 0.63) with a zirconia ferrule (OD = 2.5 mm) to the split mating sleeve on the optrode holder. Push the patch cord into the sleeve until the tip of the ferrule contacts the back of the glass pipette. The patch cord delivers the light (wavelength = 465 nm) from LED. The LED light is controlled by a single-channel LED driver.</li>
	<li>Remove the silicone adhesive over the craniotomy. Put warmed saline (38 &#176;C) on the brain surface periodically to prevent the brain surface from drying during the recording session. If necessary, remove the dura with sharp tweezers.</li>
	<li>Insert the glass electrode into the brain tissue with a manipulator. Monitor the electrode resistance when it is inserted. Replace the electrode when the resistance is lower than expected because the tip of the electrode may be damaged. At the depth of interest, search and isolate the single unit spikes by adjusting the depth in 2 to 5 &#181;m steps.</li>
	<li>After the single unit is isolated, deliver the LED light and observe the response to the light stimuli.</li>
	<li>Record the neural activities after the identification of the cell types. Perform the recordings for 2&#8211;3 h. If the recording is made on consecutive days (maximum of 2 days), cover the craniotomy with silicone adhesive. When the recording is finished, and the animal is sacrificed, retrieve the head post and put it in acetone to wash and reuse it.</li>
</ol>

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The authors have nothing to disclose.

Optogenetics has become a powerful tool in neuroscience. It has been utilized for identifying specific neuron types in vivo as well as manipulating the activities of specific neuronal pathways. The clarification of the neural activities of different neuronal types promotes the understanding of the mechanism of the neural circuits. Here, we demonstrated a method to deliver the light to the recording site through a glass electrode in the IC of awake VGAT-ChR2 mice.
There are several cri...

The central nervous system consists of various types of neurons, which have different functions. How these different types of neurons work within the neural circuit is one of the major concerns in neuroscience. However, in many brain regions, it has been impossible to distinguish the neuronal types in in vivo recordings of electrical activities because there is no clear difference in the electrical spike signal itself, with some exceptions. Recent advances in optogenetics have made a breakthrough1,2. Using transgenic animals in which light-sensitive opsin (e.g., channelrhodopsin-2) is expressed in specific neuronal types, it became possible to distinguish the neuronal types efficiently in in vivo recordings3,4,5,6. In these animals, the neurons with light-sensitive opsin are excited by giving light stimuli during the electrical recordings, but other neurons are not. The opsin-positive neurons, therefore, are easily distinguished from other neuron types by their responses to light.
In optogenetics experiments, it is critical to deliver the light to the recording site. As a non-invasive method, the light is often directed from the brain&#39;s surface. However, because the light&#39;s strength reduces as it goes through brain tissue, it is hard to stimulate 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. Electrophysiological experiments have often been performed on anesthetized animals because the body movement causes noise in the recordings. As is well documented, however, anesthesia is known to change the neural responses7,8,9,10. Thus, it is necessary to use awake animals in order to study neural responses without the artificial effects of anesthesia. Unlike the experiments with anesthetized animals, the electrophysiological recordings are performed after the recovery from surgery in the experiments with awake animals. During the interval between the surgery and the recordings, the tissue exudate often accumulates on the brain surface and makes the optical transparency of the brain surface low.
Here, we describe a method to record single-unit recordings 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 deep brain regions. This custom-made optrode system consists of accessible and inexpensive materials and is easy to assemble.

<table><tbody><tr><td>Electrode holder</td><td>Molecular Device</td><td>1-HL-U</td><td>pipette holder for microelectrode amplifier</td></tr><tr><td>Ceramic split mating sleeve</td><td>Thorlabs</td><td>ADAF1</td><td>f2.5 mm ferrule</td></tr><tr><td>Circuit board spacer</td><td>Teishin Denki</td><td>SPA-320</td><td>f8.0 mm, 20.0 mm long</td></tr><tr><td>Stereotaxic frame for mice</td><td>Narishige</td><td>SR-6M-HT</td><td>Stereotaxic instruments for mice</td></tr><tr><td>Manipulator</td><td>Narishige</td><td>NA</td><td>Manual manipulator</td></tr><tr><td>Superbond</td><td>Sun Medical</td><td>M: 204610557</td><td>Dental adhesive resin cement</td></tr><tr><td>Form2</td><td>Formlabs</td><td>NA</td><td>3D printer</td></tr><tr><td>Kwik-Sil</td><td>WPI</td><td>KWIK-SIL</td><td>Low toxicity silicone adhesive</td></tr><tr><td>Borosilicate glass capillaries</td><td>Narishige</td><td>GD-1.5</td><td>OD 1.5 mm, ID 0.9 mm, 90.0 mm long</td></tr><tr><td>Fiber-optic patch cord</td><td>Doric Lenses</td><td>MFP_960/1000/2200-0.63_1m_FCM-ZF2.5</td><td>Monofiberoptic patchcord, OD, 2.5 mm, core = 960 mm, cladding = 1000 mm, NA = 0.63</td></tr><tr><td>Connectrized LED</td><td>Doric Lenses</td><td>LEDC-1B_FC</td><td>Central wave length = 465 nm, output power = 45 mW (Core 960 mm 0.63 NA )</td></tr><tr><td>LED driver</td><td>Doric Lenses</td><td>LEDRV_1CH_1000</td><td>1 ch LED driver, maximum output = 1000 mA</td></tr><tr><td>Electrode puller</td><td>Narishige</td><td>PB-7</td><td>Dual-stage glass micropipette puller</td></tr><tr><td>Borosilicate glass capillary</td><td>Narishige</td><td>GD-1.5</td><td>Bolosilicate glass capillary, OD, 1.5mm, ID, 0.9 mm, 90.0 mm long</td></tr><tr><td>GENTACIN</td><td>MSD CO., Ltd</td><td>185711173</td><td>Antibiotic ointment</td></tr><tr><td>Terramycin&amp;reg;-LA</td><td>Zoetis</td><td>G 333</td><td>Oxytetracycline</td></tr><tr><td>Tg(Slc32a1- COP4*H134R/EYFP)8Gfng/J</td><td>Jackson Labs</td><td>#14548</td><td>VGAT-ChR2 mice</td></tr><tr><td>Multiclamp 700B</td><td>Molecular Devices</td><td>2500-0157</td><td>Microelectrode amplifier</td></tr></tbody></table>

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.

In Figure 2, we examined the effects of the tip size and the length of the glass pipettes on the light power at the tip of the pipettes (Figure 2A-B). The light power was measured by an optical power meter placed 1 mm away from the tip. The full length was set to 50 &#177; 2 mm when the tip size varied, while the tip size was set to 2.5 &#181;m when the full length varied. The shank of the glass pipette was set t...

Watch this Scientific Journal Video about Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice at JoVE.com

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

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 ...

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6.7K Views.  Kanazawa Medical University. This work introduces a method to perform an optogenetic single-unit recording reliably from an awake mouse using a custom-made glass optrode.

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

Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack conspicuous morphological features or show a low population density. This applies particularly to retinal amacrine cells, an exceptionally multiform class of interneurons that comprise roughly 30 subtypes in mammals1. Though being a crucial part of the visual processing by shaping the retinal output2, most of these subtypes have not been studied up to now in a functional context because encountering these cells with a recording electrode is a rare event.
Recently, a multitude of transgenic mouse lines is available that express fluorescent markers like green fluorescent protein (GFP) under the control of promoters for membrane receptors or enzymes that are specific to only a subset of neurons in a given tissue3,4. These pre-labeled cells are therefore accessible to directed microelectrode targeting under microscopic control, permitting the systematic study of their physiological properties in situ. However, excitation of fluorescent markers is accompanied by the risk of phototoxicity for the living tissue. In the retina, this approach is additionally hampered by the problem that excitation light causes appropriate stimulation of the photoreceptors, thus inflicting photopigment bleaching and transferring the retinal circuits into a light-adapted condition. These drawbacks are overcome by using infrared excitation delivered by a mode-locked laser in short pulses of the femtosecond range. Two-photon excitation provides energy sufficient for fluorophore excitation and at the same time restricts the excitation to a small tissue volume minimizing the hazards of photodamage5. Also, it leaves the retina responsive to visual stimuli since infrared light (&gt;850 nm) is only poorly absorbed by photopigments6.
In this article we demonstrate the use of a transgenic mouse retina to attain electrophysiological in situ recordings from GFP-expressing cells that are visually targeted by two-photon excitation. The retina is prepared and maintained in darkness and can be subjected to optical stimuli which are projected through the condenser of the microscope (Figure 1). Patch-clamp recording of light responses can be combined with dye filling to reveal the morphology and to check for gap junction-mediated dye coupling to neighboring cells, so that the target cell can by studied on different experimental levels.

This article depicts the recording of individual cells from fluorescently tagged neuronal populations in the intact mouse retina. By using two-photon infrared excitation transgenetically labeled cells were targeted for patch-clamp recording to study their light responses, receptive field properties, and morphology.

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack ...

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 ...

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits&#160;in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.

We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations&#160;in vitro.

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the ...

Optogenetic Manipulation of Neuronal Activity to Modulate Behavior in Freely Moving Mice

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of single neurons and region-specific transmitter release, up to whole neuronal circuitries in the central nervous system, and allows the direct measurement of behavioral outcomes. Neurons express optogenetic tools via an injection of viral vectors carrying the DNA of choice, such as Channelrhodopsin2 (ChR2). Light is brought into specific brain regions via chronic optical implants that terminate directly above the target region. After two weeks of recovery and proper tool-expression, mice can be repeatedly used for behavioral tests with optogenetic stimulation of the neurons of interest.
Optogenetic modulation has a high temporal and spatial resolution that can be accomplished with high cell specificity, compared to the commonly used methods such as chemical or electrical stimulation. The light does not harm neuronal tissue and can therefore be used for long-term experiments as well as for multiple behavioral experiments in one mouse. The possibilities of optogenetic tools are nearly unlimited and enable the activation or silencing of whole neurons, or even the manipulation of a specific receptor type by light.
The results of such behavioral experiments with integrated optogenetic stimulation directly visualizes changes in behavior caused by the manipulation. The behavior of the same animal without light stimulation as a baseline is a good control for induced changes. This allows a detailed overview of neuronal types or neurotransmitter systems involved in specific behaviors, such as anxiety. The plasticity of neuronal networks can also be investigated in great detail through long-term stimulation or behavioral observations after optical stimulation. Optogenetics will help to enlighten neuronal signaling in several kinds of neurological diseases.

With optogenetic manipulation of specific neuronal populations or brain regions, behavior can be modified with high temporal and spatial resolution in freely moving animals. By using different optogenetic tools in combination with chronically implanted optical fibers, a variety of neuronal modulations and behavioral testing can be performed.

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of ...

Behavior

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 ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. Few studies have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem, which contains second order neurons of the auditory pathway, is a potential site for gene transfer. In this protocol, a technique for direct and maximal exposure of the murine DCN via a posterior fossa approach is demonstrated. This approach allows for either acute or survival surgery. Following direct visualization of the DCN, a host of experiments are possible, including injection of opsins into the cochlear nucleus and subsequent stimulation by an optical fiber coupled to a blue light laser.&#160;Other neurophysiology experiments, such as electrical stimulation and neural injector tracings are also feasible.&#160;The level of visualization and the duration of stimulation achievable make this approach applicable to a wide range of experiments.

The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model.

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. ...

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 ...

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.

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 ...

Installation of Optical Windows in the Brain of Transgenic Mice to Image Neural Activities

Source: Lin, X., et al. <a href="https://app.jove.com/v/56297">Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice.</a> J. Vis. Exp. (2019).
This video demonstrates the brain&#39;s neural activity recording through an optical window in transgenic mice. The mice express a fluorescent protein that enables the detection of neural activities. A small portion of the skull is carefully removed to install an optical window, and the skull is then covered with black cement to block light before imaging.

Instalacja okien optycznych w mózgu myszy transgenicznych w celu zobrazowania aktywności neuronalnej

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice

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Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

Optogenetic Manipulation of Neuronal Activity to Modulate Behavior in Freely Moving Mice

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

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

Optrode Array for Simultaneous Optogenetic Modulation and Electrical Neural Recording

Instalacja okien optycznych w mózgu myszy transgenicznych w celu zobrazowania aktywności neuronalnej