This work was supported by the Korea Institute of Science and Technology Institutional Program (Projects No. 2E26190 and 2E26170) and the Human Frontier Science Program (RGY0089/2012).

All methods described have been approved by the Animal Care and Use Committee of the Korea Institute of Science and Technology.
1. Preparing the Micro-drive and Electrode
<ol>
	<li>Assembling the micro-drive.
	<ol>
		<li>Print the parts of the micro-drive (slider, body, and shell)14 using a high-resolution 3D printer. Make sure the parts have no defects.</li>
		<li>Fix the slider into the micro-drive body with a screw (size 000-120x1/4).</li>
		<li>Solder a nut (size 000-120x5/64 Hex) onto the tip of the screw.</li>
		<li>Rotate the screw clockwise to move the slider up or counter-clockwise to move it down.</li>
		<li>Fix the shell to the body with a screw (size 000-120x1/4).</li>
	</ol>
	</li>
	<li>Mounting the probe on the micro-drive.
	<ol>
		<li>Fix the micro-drive in a horizontal position under a binocular microscope.</li>
		<li>Use a flat alligator clip with rubber padding to grab the flex cable of the silicon probe, while carefully detaching the probe from the shipment enclosure with forceps. Fix the alligator clip to a manipulator.</li>
		<li>Using the manipulator, lay the silicon probe on the micro-drive slider, parallel to the direction of movement.</li>
		<li>Apply a drop of dental cement (room temperature-curing acrylic dental repair material) to fix the probe to the slider. Correct the probe position if it has shifted. Glue is not recommended as it cures very fast and makes it difficult to readjust the electrode position.</li>
		<li>Fix the connector of the probe to the C-holder (connector holder) with dental cement.</li>
	</ol>
	</li>
	<li>Cleaning and putting dye on the probe.
	<ol>
		<li>Fix the Microdrive/probe assemblage on the probe-cleaning device. The device is equipped with 2 small rotating foam sponges (non-sterile swabs). Adjust the gap by the manipulator.</li>
		<li>Soak the sponges with detergent.</li>
		<li>Slowly move the probe up and down between the sponges, allowing a gentle touch. Monitor the cleaning process under a microscope.</li>
		<li>Rinse the probe with distilled water. Dip the probe in alcohol for sterilization.</li>
		<li>Apply Dil (lipophilic fluorescent dye) on the back of the shanks of the probe, using a foam swab. This will allow in a later stage the visualization of shank locations in the brain.</li>
	</ol>
	</li>
	<li>Assembling the hat
	<ol>
		<li>Print the parts of the hat (head-plate, connector holder, and cap)14 using a 3D-printer. The 3 parts fit together to form a sealed enclosure.</li>
		<li>Insert nuts (size M2 and 00-90x1/4) in the slots of the head-plate and fix with glue and dental cement.</li>
		<li>Insert a washer in the slot of the cap and fix with dental cement.</li>
	</ol>
	</li>
</ol>
2. Surgery for Fixation of the Hat on the Skull
All tools used for surgery are sterilized beforehand by autoclaving. A dry heat beads device is used forto sterilize tools that become contaminated and need to be sterilized during the surgery.
<ol>
	<li>Set the level of isoflurane to 4% for the initiation of anesthesia. Put the mouse in the anesthesia chamber for 5 min.</li>
	<li>Install the mouse in a stereotaxic apparatus.</li>
	<li>Reduce the isoflurane level to 1.5-2%. Adjust the level during the surgery according to animal state, breathing rate, and body temperature20.</li>
	<li>Apply eye ointment on the eyes.</li>
	<li>Shave the scalp and clean the head of the animal with antiseptic (iodopovidone). Maintain aseptic conditions during all steps of the surgery.</li>
	<li>Inject bupivacaine (1 mg/kg) under the scalp. Cut and remove part of the parietal skin of the mouse head using fine scissors to expose the skull at its edges. Use saline and a hemostatic sponge to clean and control the bleeding during the surgery.</li>
	<li>Remove the periosteum using a scraper tool.</li>
	<li>Find the reference points on the skull21: bregma, lambda, coronal, and sagittal suture. Adjust the head&#39;s angle along the sagittal axis such that the Bregma and lambda points are at the same height.</li>
	<li>Drill two holes (~0.5 mm diameter) in the skull for the reference and ground electrodes. The holes should be approximately 1 mm caudal and 1 mm lateral of the Lambda.</li>
	<li>Insert the ground and reference electrodes (size 000-120x1/16 miniature stainless-steel screws wire-coupled to pin connectors).</li>
	<li>Apply ultraviolet (UV) light bonding dentin activator on the skull and then apply UV light for 45-60 s.</li>
	<li>Apply a layer of dental cement on the edges of the skull. Avoid spreading cement on the skin and hair of the mice.</li>
	<li>Fix the head-plate to a stereotaxic manipulator and position it above the skull. Slowly lower the head-plate until it slightly touches the skull, and apply dental cement at the junction with the skull. Let the dental cement cure for 15 min.</li>
	<li>Remove the anesthesia. Fix a connector-holder and a cap to the head-plate. Put the mouse in its cage after giving a sub-cutaneous injection of ketaprofen 5 mg/kg.</li>
	<li>Give sub-cutaneous injections of ketaprofen 5 mg/kg for the two next days and monitor carefully for any sign of pain. Mice typically wake up from anesthesia within 20-40 min. The hat implant is relatively light (3.34g), such that mice have no trouble to lift their head, run in mazes and climb on the edges of their home cage.</li>
</ol>
3. Behavioral Training
<ol>
	<li>After a post-surgery recovery period of 7 days, start the restriction of water at 1 mL per day.</li>
	<li>To make the treadmill belt, cut a piece of velvet fabric (5 cm by 1-2 m) and fix small objects on it using hot glue. Attach erected objects on the edges of the belt in order not to interfere with the motion of the mouse.</li>
	<li>Fix the belt on the wheels of the treadmill by suturing together the two ends.</li>
	<li>Install the mouse head-fixed in the treadmill by inserting and tightening the two screws of the head-plate into the slots of the head-fixation plate.</li>
	<li>Start training the mouse to run head-restrained on the treadmill for the water reward. Deliver the water reward through a lick port. Initially, put the mouse on the treadmill for periods of 10 min, 3 times per day.</li>
	<li>As the mouse gets used to the head-fixation and starts to move the belt (typically after ~3 days), increase the training session duration to 30 min. After 2-3 weeks, some mice run more than 100 trials in 30 min (a trial being a full rotation of the belt).</li>
	<li>Choose the mice showing the best behavioral performances for the recording experiments.</li>
</ol>
4. Implantation of the Silicon Probe
<ol>
	<li>Put the mouse under anesthesia.</li>
	<li>Install the mouse in the stereotaxic apparatus by fixing the head-plate. Clean the skull surface with saline.</li>
	<li>Find stereotaxic markers: bregma, lambda, coronal, and sagittal suture. Measure the distance to the point of insertion and mark it.</li>
	<li>Drill the bone carefully until it becomes thin and transparent. Moisten and clean with saline while drilling.</li>
	<li>Carefully remove the thinned bone and the Dura matter using precision forceps. Keep the brain surface wet with saline all the time.</li>
	<li>Fix the Microdrive/silicon probe assemblage to a stereotaxic manipulator. Bring the silicon probe just above the craniotomy. Screw the silicon probe connector holder to the head-plate.</li>
	<li>Connect the recording amplifier and ground/reference electrodes. Shield the mouse with aluminum foil to protect from electro-magnetic noise. Start the recording system to monitor the electrical activity of the brain.</li>
	<li>Slowly insert the silicon probe into the brain using the micromanipulator. Continuously check the electrical signals, the manipulator travelled distance, and the shanks of the probe (make sure they are penetrating in the brain). Unit activity is visible in the cortex while the white matter underneath is relatively silent. Unit activity reappears when the shanks touch the pyramidal layer of the hippocampus. From this point, retract the silicon probe 200 &#181;m (use the micro-drive the next day to bring the shank back into the pyramidal layer).</li>
	<li>Cover the surface of the brain with a mixture of bone wax and mineral oil.</li>
	<li>Fix the micro-drive to the head-plate using dental cement. Let the dental cement cure for 15 min. Then detach the micro-drive from the stereotaxic manipulator and put the hat cap back on.</li>
	<li>Put the mouse back in its cage and give a sub-cutaneous injection of ketaprofen 5 mg/kg. Check for any sign of pain. This surgery is much less invasive than the first one and mice are typically active within 45 min after they wake-up from the anesthesia. However, let the mouse recover an entire day as the silicon probe needs to stabilize in the brain.</li>
</ol>
5. Recording
<ol>
	<li>Install the mouse head-fixed on the treadmill. Remove the hat cap.</li>
	<li>Connect the recording amplifier and start the recording.</li>
	<li>On the first day, use the micro-drive to move the silicon probe into the pyramidal layer of the hippocampus. Each counter-clockwise turn of the screw moves the shanks 200 &#181;m deeper. Adjust the shank position slowly (~20-50 &#181;m at a time) until ripple oscillations13 and unit activity appear.</li>
	<li>On the next days, tune the shank position and wait ~1 h before recording the hippocampal activity while the mouse runs on the belt. To maintain good recording signal quality for several days, remove hard objects and low ceiling from the mouse&#39;s cage to minimize the chance the hat bumps into hard surfaces.</li>
</ol>
6. Recovering the probe
<ol>
	<li>Put the mouse under anesthesia.</li>
	<li>Install the mouse in the stereotaxic apparatus by fixing the head-plate. Remove the hat cap.</li>
	<li>Bring the stereotaxic manipulator just above the micro-drive. Fix the micro-drive to the manipulator. Unscrew the connector holder from the head-plate. Remove the screw connecting the shell and body of the micro-drive.</li>
	<li>Under binocular microscope supervision, slowly pull up the micro-drive/silicon probe assemblage with the stereotaxic manipulator, leaving the shell part behind.</li>
	<li>Clean the silicon probe with the cleaning device.</li>
</ol>

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The authors have nothing to disclose.

Chronic recording of neuronal activity is critical for understanding neural processes such as hippocampal place fields. Our approach to perform chronic silicon probe implantantation distinguishes itself from other methods7,18,19,20 by the fact that it is relatively simple to recover the electrode package at the end of the experiment. While place cells are typically studied in freely-moving cond...

Silicon probes present several advantages for electrophysiological recordings, including the fact that they are designed with sharp profiles minimizing tissue damage and that they present a precise layout of densely packed recording sites1,2,3,4. They are used to study various systems in different species, including humans3,5,6, with diverse approaches1,7. Yet, their recurrent use is still relatively limited because of their cost, fragility, and the fact that convenient methods for chronic experiments are lacking8. Recent advances in 3D printing technology have made possible the custom designing of devices such as micro-drives and head-plates to allow an easier handling of these delicate electrodes. In a first step, we will describe how to build and use a set of tools that we have developed for the implantation of chronic silicon probes14.
While place cells are typically studied using freely-moving animals running in mazes, recently they were also investigated in virtual environments15 and in treadmill apparatii9 (Figure 1A). These experimental methods offer the advantage that animals can be head-restrained, making the use of 2-photon microscope15, patch-clamp16, and optrode9,10,11 techniques easier, in addition to providing enhanced control on animal behavior and environmental cues12. In a second step, we will present the procedures for training mice and recording place cell activity in a treadmill apparatus.

<table>
	<tbody>
		<tr>
			<td>Silicon Probe</td>
			<td>Neuronexus</td>
			<td>Buzsabi32</td>
			<td>Recording electrode</td>
		</tr>
		<tr>
			<td>Recording system</td>
			<td>Intantech</td>
			<td>RHD2132/RHD2000</td>
			<td></td>
		</tr>
		<tr>
			<td>3D printer</td>
			<td>Asiga</td>
			<td>Pico Plus 27</td>
			<td>High resolution printer for micro-drive</td>
		</tr>
		<tr>
			<td>3D printer</td>
			<td>Stratasys</td>
			<td>Mojo</td>
			<td>Lower resolution printer for hat components</td>
		</tr>
		<tr>
			<td>Stereotaxic apparatus</td>
			<td>Kopf</td>
			<td>Model 963</td>
			<td></td>
		</tr>
		<tr>
			<td>Binocular microscope</td>
			<td>Leica</td>
			<td>M60</td>
			<td></td>
		</tr>
		<tr>
			<td>Treadmill apparatus</td>
			<td></td>
			<td></td>
			<td>We build them</td>
		</tr>
	</tbody>
</table>

implantation of chronic silicon probes and recording of hippocampal place cells in an enriched treadmill apparatus

An important requisite for understanding brain function is the identification of behavior and cell activity correlates. Silicon probes are advanced electrodes for large-scale electrophysiological recording of neuronal activity, but the procedures for their chronic implantation are still underdeveloped. The activity of hippocampal place cells is known to correlate with an animal&#39;s position in the environment, but the underlying mechanisms are still unclear. To investigate place cells, here we describe a set of techniques which range from the fabrication of devices for chronic silicon probe implants to the monitoring of place field activity in a cue-enriched treadmill apparatus. A micro-drive and a hat are built by fitting and fastening together 3D-printed plastic parts. A silicon probe is mounted on the micro-drive, cleaned, and coated with dye. A first surgery is performed to fix the hat on the skull of a mouse. Small landmarks are fabricated and attached to the belt of a treadmill. The mouse is trained to run head-fixed on the treadmill. A second surgery is performed to implant the silicon probe in the hippocampus, following which broadband electrophysiological signals are recorded. Finally, the silicon probe is recovered and cleaned for reuse. The analysis of place cell activity in the treadmill reveals a diversity of place field mechanisms, outlining the benefit of the approach.

A mouse was first trained to run on a two-meter long belt devoid of cues (Figure 1C). Following electrode implantation, a new belt of the same length but presenting 3 pairs of cues was installed on the treadmill, in order to generate allocentric spatial representations12,14. Broadband signals were recorded at a sampling rate of 30,000 Hz, using a 250-channel recording system (amplifier board with USB ...

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Implantation of Chronic Silicon Probes and Recording of Hippocampal Place Cells in an Enriched Treadmill Apparatus

We describe the diverse steps to implant chronic silicon probes and to record place cells in mice that are running head-fixed on a cue-enriched treadmill apparatus.

An important requisite for understanding brain function is the identification of behavior and cell activity correlates. Silicon probes are advanced ...

implantation-chronic-silicon-probes-recording-hippocampal-place-cells

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12.6K Views.  Korea Institute of Science and Technology. We describe the diverse steps to implant chronic silicon probes and to record place cells in mice that are running head-fixed on a cue-enriched treadmill apparatus.

Video: Implantation of Chronic Silicon Probes and Recording of Hippocampal Place Cells in an Enriched Treadmill Apparatus

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Recording Large-scale Neuronal Ensembles with Silicon Probes in the Anesthetized Rat

Large scale electrophysiological recordings from neuronal ensembles offer the opportunity to investigate how the brain orchestrates the wide variety of behaviors from the spiking activity of its neurons. One of the most effective methods to monitor spiking activity from a large number of neurons in multiple local neuronal circuits simultaneously is by using silicon electrode arrays1-3.
Action potentials produce large transmembrane voltage changes in the vicinity of cell somata. These output signals can be measured by placing a conductor in close proximity of a neuron. If there are many active (spiking) neurons in the vicinity of the tip, the electrode records combined signal from all of them, where contribution of a single neuron is weighted by its 'electrical distance'. Silicon probes are ideal recording electrodes to monitor multiple neurons because of a large number of recording sites (+64) and a small volume. Furthermore, multiple sites can be arranged over a distance of millimeters, thus allowing for the simultaneous recordings of neuronal activity in the various cortical layers or in multiple cortical columns (Fig. 1). Importantly, the geometrically precise distribution of the recording sites also allows for the determination of the spatial relationship of the isolated single neurons4. Here, we describe an acute, large-scale neuronal recording from the left and right forelimb somatosensory cortex simultaneously in an anesthetized rat with silicon probes (Fig. 2).

Extracellular recordings of neuronal activity using silicon probes in the anesthetized rat will be described. This technique allows information to be obtained across multiple brain areas from more than 100 neurons simultaneously. It provides information with single cell resolution about neuronal ensembles dynamics in multiple local circuits.

Large scale electrophysiological recordings from neuronal ensembles offer the opportunity to investigate how the brain orchestrates the wide variety of ...

Surgical Implantation of Chronic Neural Electrodes for Recording Single Unit Activity and Electrocorticographic Signals

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery. Here we provide useful information for surgeons who are learning the process of implanting electrode systems. We demonstrate the implantation procedure of both a penetrating and a surface electrode. The surgical process is described from start to finish, including detailed descriptions of each step throughout the procedure. It should also be noted that this video guide is focused towards procedures conducted in rodent models and other small animal models. Modifications of the described procedures are feasible for other animal models.

We provide useful information for surgeons who are learning the process of implanting chronic neural recording electrodes. Techniques for both penetrating and surface electrode systems are described in a rodent animal model.

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery.  Here we provide useful information for ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

Design and Assembly of an Ultra-light Motorized Microdrive for Chronic Neural Recordings in Small Animals

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of neural circuits underlying a variety of natural behaviors, including navigation1 , decision making 2,3, and the generation of complex motor sequences4,5,6. Advances in precision machining has allowed for the fabrication of light-weight devices suitable for chronic recordings in small animals, such as mice and songbirds. The ability to adjust the electrode position with small remotely controlled motors has further increased the recording yield in various behavioral contexts by reducing animal handling.6,7
 Here we describe a protocol to build an ultra-light motorized microdrive for long-term chronic recordings in small animals. Our design evolved from an earlier published version7, and has been adapted for ease-of use and cost-effectiveness to be more practical and accessible to a wide array of researchers. This proven design 8,9,10,11 allows for fine, remote positioning of electrodes over a range of ~ 5 mm and weighs less than 750 mg when fully assembled. We present the complete protocol for how to build and assemble these drives, including 3D CAD drawings for all custom microdrive components.

The design, fabrication and assembly of an ultra-light motorized microdrive is described. The device provides a cost-effective and easy-to-use solution for chronic recordings of single units in small behaving animals.

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

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

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

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