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 critical steps in the method described. First, it is a requirement to use animals with a sufficient and specific expression of opsins in the target neurons. To identify the responses to light with the extracellular recordings, it is necessary that the activation of the opsin evokes the spikes in the target neurons. In some transgenic strains, it is possible that the transgene expression level in neurons is low in young animals14, and there might be a minimum age for the experiment. Further, it may be necessary to perform some additional experiments to confirm if the responses to the light stimuli are specific to the target neuron types (e.g., immunohistochemistry). When the target neuron is excitatory, it would be necessary to distinguish the directly evoked responses from the synaptically evoked responses, which could be done by thresholding the latency of the response. Second, it is critical to ensure the brain tissue is as clean as possible. A maintenance of cleanliness is especially required in order to prevent any bleeding during the craniotomy and the recordings. Also, the surface of the brain should be prevented from drying after the craniotomy by covering the surface tightly with silicone adhesive.
To deliver the light efficiently from the electrode tip, it is critical to reduce any loss in the optical path. In our configuration of the optrode, the light power drops tenfold through the electrode holder: the light power is 45 mW at the end of the fiber-optic patch cord, while it is 3&#8211;5 mW at the tip of the glass pipette. As shown in the results, the tip size and the full length did not affect the light power at the tip (Figure 2). These results indicated that the light attenuation was fairly stable regardless of the configuration of the glass pipette. This light attenuation did not matter in the recording of IC neurons in VGAT-ChR2 mice because we have confirmed that it was possible to record light activated spikes in the ventral edge (2 mm depth) of the IC, although it may matter in other recording configurations. It would be hard to reduce the light attenuation dramatically, but there may be possibilities for improvement. First, it may be effective to use quartz instead of a borosilicate glass as an electrode pipette. It is known that quartz has better light conductivity, so that it may reduce the light loss through the electrode15. Also, it was reported that the polish of the end of the pipette reduced light loss at the contact surface between the pipette and patch cord15. If the improvement in the light attenuation is not enough, it would be effective to use a laser instead of LED, although laser costs more than LED.
One of the advantages of the glass pipette electrode is that it is possible to perform histological procedures by adding tracers or vectors to the filling solution. The histological procedures will add a great deal of useful information (the location of the recording, the morphology of the recorded neuron, etc.) to the electrophysiological data. Also, it was reported that it is possible to perform pharmacological surveys using the glass electrode with blockers in the filling solution15. In contrast to these advantages, the limitation of the glass pipette electrode is that it cannot record from multiple isolated units. To attain multiple recordings, it may be possible to put a tetrode wire into the glass pipette instead of a single Ag-Cl coated wire. In this case, it would be necessary to use a pipette with a larger tip size (30&#8211;60 &#181;m, the typical tip size of a tetrode) and let the end of the tetrode out from the tip of the pipette.
While previously reported optrodes often required complex fabrication16,17, Magee and his colleagues have developed an optrode using standard glass pipettes as with the optrode we designed18. In their method, they insert a thin optical fiber into the pipette to deliver the light. To make the fiber thin enough to insert the shank of the pipette, they etched 125 &#181;m cladding fiber with a 9 &#181;m core to a diameter of 8&#8211;10 &#181;m at the tip. The advantage of this method is that it would apply even for the bent pipette that is often used for iontophoresis. However, it might be hard to use LED as a light source because standard commercial LEDs would not achieve sufficient light power via a 9 &#181;m core fiber.

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 border="0" cellpadding="0" cellspacing="0" style="width:971px;" width="971">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:247px;">Electrode holder</td>
			<td style="width:140px;">Molecular Device</td>
			<td style="width:393px;">1-HL-U</td>
			<td style="width:191px;">pipette holder for microelectrode amplifier</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Ceramic split mating sleeve</td>
			<td>Thorlabs</td>
			<td>ADAF1</td>
			<td>f2.5 mm ferrule</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Circuit board spacer</td>
			<td>Teishin Denki</td>
			<td>SPA-320</td>
			<td>f8.0 mm, 20.0 mm long</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Stereotaxic frame for mice</td>
			<td>Narishige</td>
			<td>SR-6M-HT</td>
			<td>Stereotaxic instruments for mice</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Manipulator</td>
			<td>Narishige</td>
			<td>NA</td>
			<td>Manual manipulator</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Superbond</td>
			<td>Sun Medical</td>
			<td>M: 204610557</td>
			<td>Dental adhesive resin cement</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Form2</td>
			<td>Formlabs</td>
			<td>NA</td>
			<td>3D printer</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Kwik-Sil</td>
			<td>WPI</td>
			<td>KWIK-SIL</td>
			<td>Low toxicity silicone adhesive</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">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 height="40">
			<td height="40" style="height:40px;">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 height="40">
			<td height="40" style="height:40px;">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 height="40">
			<td height="40" style="height:40px;">LED driver</td>
			<td>Doric Lenses</td>
			<td>LEDRV_1CH_1000</td>
			<td>1 ch LED driver, maximum output = 1000 mA</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Electrode puller</td>
			<td>Narishige</td>
			<td>PB-7</td>
			<td>Dual-stage glass micropipette puller</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:247px;">Borosilicate glass capillary</td>
			<td style="width:140px;">Narishige</td>
			<td style="width:393px;">GD-1.5</td>
			<td style="width:191px;">Bolosilicate glass capillary, OD, 1.5mm, ID, 0.9 mm, 90.0 mm long</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">GENTACIN</td>
			<td>MSD CO., Ltd</td>
			<td>185711173</td>
			<td>Antibiotic ointment</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Terramycin&#174;-LA</td>
			<td>Zoetis</td>
			<td>G 333</td>
			<td>Oxytetracycline</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Tg(Slc32a1- COP4*H134R/EYFP)8Gfng/J</td>
			<td>Jackson Labs</td>
			<td>#14548</td>
			<td>VGAT-ChR2 mice</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">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 to 8 mm. The light power at the tip ranged from 3&#8211;5 mW (mean &#177; SD; Figure 2A: 3.77 &#177; 0.29 mW, n = 20; Figure 2B: 4.24 &#177; 0.31 mW, n = 21). The tip size (Figure 2A) of the pipettes was hardly correlated with the light power (R = 0.1555). The length of the pipette was only slightly correlated with the light power in a negative manner (R = -0.2054). These results suggest that the tip and length of the pipettes rarely affect the light power at the tip. We also checked if the amount of volume load of the solution (10 mM PBS) in the glass pipettes affects the light power (Figure 2C). The volume of the solution was assessed by the length of the solution in the pipette. The data were recorded from 4 pipettes (tip size: 0.25 &#181;m, the full length: 50 &#177; 2 mm, the length of the shank: 8 mm). We found that varying the volume of the solution in the pipette did not affect the light power (Figure 2C).
In Figure 3, we show the spike responses recorded in the inferior colliculus (IC) of awake VGAT-ChR2 mice (2&#8211;4 months old). The IC is an auditory center in the midbrain and consists of glutamatergic and GABAergic neurons12,13, both of which respond to sound3,4. In the IC of VGAT-ChR2 mice, the GABAergic neurons specifically express ChR24. In previous work, it was shown that the GABAergic and glutamatergic neurons were excited or suppressed, respectively, when they were given light stimuli3,4. In the awake mice, we found both types of neurons (Figure 3A-B). When the light stimulus is delivered through the glass electrode, it evoked spike responses in some neurons (Figure 3A). In contrast, in other neurons, the light stimuli suppressed the spike responses evoked by sound stimuli (Figure 3B). These results show that a glass optrode can record the well-isolated single unit activity, and reliably stimulate the recorded neurons by light.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57781/57781fig1.jpg" /> 
Figure 1: Optrode holder and recording accessories. Panels (A) and (B) show the optrode holder. Panel (A) shows the part structure of the optrode holder. (1) This is a ceramic split mating sleeve. (2) This is a stainless-steel tube. (3) This is a barrel of the holder. (4) This is a cone washer. (5) This is a polycarbonate cap. Panel (B) shows the assembled optrode holder. Panels (C) and (D) show the custom-built head post. Panel (C) shows a side view of the head post. Panel (D) shows an oblique view of the head post. The two head posts (right) are made from a circuit board spacer (left). Panels (E), (F), and (G) show the custom-made resin half tube. Panels (E) and (F) show dimensional drawings of the resin half tube. Panel (E) shows a front view of the resin half tube. Panel (F) shows a side view of the resin half tube. Panel (G) shows a photo image of the half tube. The numbers in panels (E) and (F) indicate the drawing dimensions in mm. <a href="/files/ftp_upload/57781/57781fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57781/57781fig2.jpg" /> 
Figure 2: The effect of the configuration of the glass pipettes on the light power at the tip. (A) The tip&#39;s size was plotted against the light power. The full length of the pipette was set to 50 &#177; 2 mm. The length of the shank was set to 8 mm. (B) The length of the pipette was plotted against the light power. The tip size was set to 0.25 &#181;m. The length of the shank was set to 8 mm. (C) This panel shows the effect of the volume load of the solution (10 mM PBS) on the light power. The data from 4 pipettes are indicated by different symbols and lines. <a href="/files/ftp_upload/57781/57781fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57781/57781fig3.jpg" /> 
Figure 3: The light-evoked responses in the IC of awake VGAT-ChR2 mice. (A) The light stimuli (blue box, 0.03 s) evoked spike responses. (B) The light stimuli (blue box, 0.03 s) suppressed the spike responses evoked by sound (gray box). The upper 3 traces are the response to sound, and the lower 3 traces are the response when both sound and light were given. <a href="/files/ftp_upload/57781/57781fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

optogenetics-identification-neuronal-type-with-glass-optrode-awake

Nanyang Technological University

Research

JoVE Journal

Neuroscience

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

Efficient Derivation of Human Neuronal Progenitors and Neurons from Pluripotent Human Embryonic Stem Cells with Small Molecule Induction

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system (CNS) structure and circuitry in today's healthcare industry. Cell-based therapies hold great promise to restore the lost nerve tissue and function for CNS disorders. However, cell therapies based on CNS-derived neural stem cells have encountered supply restriction and difficulty to use in the clinical setting due to their limited expansion ability in culture and failing plasticity after extensive passaging1-3. Despite some beneficial outcomes, the CNS-derived human neural stem cells (hNSCs) appear to exert their therapeutic effects primarily by their non-neuronal progenies through producing trophic and neuroprotective molecules to rescue the endogenous cells1-3. Alternatively, pluripotent human embryonic stem cells (hESCs) proffer cures for a wide range of neurological disorders by supplying the diversity of human neuronal cell types in the developing CNS for regeneration1,4-7. However, how to channel the wide differentiation potential of pluripotent hESCs efficiently and predictably to a desired phenotype has been a major challenge for both developmental study and clinical translation. Conventional approaches rely on multi-lineage inclination of pluripotent cells through spontaneous germ layer differentiation, resulting in inefficient and uncontrollable lineage-commitment that is often followed by phenotypic heterogeneity and instability, hence, a high risk of tumorigenicity7-10. In addition, undefined foreign/animal biological supplements and/or feeders that have typically been used for the isolation, expansion, and differentiation of hESCs may make direct use of such cell-specialized grafts in patients problematic11-13. To overcome these obstacles, we have resolved the elements of a defined culture system necessary and sufficient for sustaining the epiblast pluripotence of hESCs, serving as a platform for de novo derivation of clinically-suitable hESCs and effectively directing such hESCs uniformly towards clinically-relevant lineages by small molecules14 (please see a schematic in Fig. 1). Retinoic acid (RA) does not induce neuronal differentiation of undifferentiated hESCs maintained on feeders1, 14. And unlike mouse ESCs, treating hESC-differentiated embryoid bodies (EBs) only slightly increases the low yield of neurons1, 14, 15. However, after screening a variety of small molecules and growth factors, we found that such defined conditions rendered retinoic acid (RA) sufficient to induce the specification of neuroectoderm direct from pluripotent hESCs that further progressed to neuroblasts that generated human neuronal progenitors and neurons in the developing CNS with high efficiency (Fig. 2). We defined conditions for induction of neuroblasts direct from pluripotent hESCs without an intervening multi-lineage embryoid body stage, enabling well-controlled efficient derivation of a large supply of human neuronal cells across the spectrum of developmental stages for cell-based therapeutics.

We have established a protocol for induction of neuroblasts direct from pluripotent human embryonic stem cells maintained under defined conditions with small molecules, which enables derivation of a large supply of human neuronal progenitors and neuronal cell types in the developing CNS for neural repair.

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Super-resolution Imaging of Neuronal Dense-core Vesicles

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and medical applications. Strengths of fluorescence include its sensitivity, specificity, and compatibility with live imaging. Unfortunately, conventional forms of fluorescence microscopy suffer from one major weakness, diffraction-limited resolution in the imaging plane, which hampers studies of structures with dimensions smaller than ~250 nm. Recently, this limitation has been overcome with the introduction of super-resolution fluorescence microscopy techniques, such as photoactivated localization microscopy (PALM). Unlike its conventional counterparts, PALM can produce images with a lateral resolution of tens of nanometers. It is thus now possible to use fluorescence, with its myriad strengths, to elucidate a spectrum of previously inaccessible attributes of cellular structure and organization.
Unfortunately, PALM is not trivial to implement, and successful strategies often must be tailored to the type of system under study. In this article, we show how to implement single-color PALM studies of vesicular structures in fixed, cultured neurons. PALM is ideally suited to the study of vesicles, which have dimensions that typically range from ~50-250 nm. Key steps in our approach include labeling neurons with photoconvertible (green to red) chimeras of vesicle cargo, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a high-resolution PALM image. We also demonstrate the efficacy of our approach by presenting exceptionally well-resolved images of dense-core vesicles (DCVs) in cultured hippocampal neurons, which refute the hypothesis that extrasynaptic trafficking of DCVs is mediated largely by DCV clusters.

We describe how to implement photoactivated localization microscopy (PALM)-based studies of vesicles in fixed, cultured neurons. Key components of our protocol include labeling vesicles with photoconvertible chimeras, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a super-resolution image.

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

In the central nervous system, the D1-alpha subtype receptor (Drd1&#945;) is the most abundant dopamine (DA) receptor, which plays a vital role in regulating neuronal growth and development. However, the mechanisms underlying Drd1&#945; receptor abnormalities mediating behavioral responses and modulating working memory function are still unclear. Using a novel RNA in situ hybridization assay, the current study identified dopamine Drd1&#945; receptor and tyrosine hydroxylase (TH) RNA expression from DA-related circuitry in the nucleus accumbens (NAc) area and substantia nigra region (SNR), respectively. Drd1&#945; expression in the NAc shows a &#34;discrete dot&#34; staining pattern. Clear sex differences in Drd1&#945; expression were observed. In contrast, TH shows a &#34;clustered&#34; staining pattern. Regarding TH expression, female rats displayed a higher signal expression per cell relative to male animals. The methods presented here provide a novel in situ hybridization technique for investigating changes in dopamine system dysfunction during the progression of central nervous system diseases.

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

Identification of dopamine D1-alpha receptor in the nucleus accumbens is critical for clarifying D1 receptor dysfunction during a central nervous system disease. We performed a novel RNA in situ hybridization assay to visualize single RNA molecules in a specific brain area.

In the central nervous system, the D1-alpha subtype receptor (Drd1&#945;) is the most abundant dopamine (DA) receptor, which plays a vital role in ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.