This research was supported by the National Natural Science Foundation of China (31871085), the Natural Science Foundation of Shanghai (21ZR1407300), the Shanghai Municipal Science and Technology Major Project (2018SHZDZX01), ZJ Lab, and Shanghai Center for Brain Science and Brain-Inspired Technology.

This protocol received approval from the Animal Care and Use Committee at Fudan University and was conducted following the guidelines and regulations designed by the National Institutes of Health Guide for Care and Use of Laboratory Animals. All possible measures were implemented to minimize the number of animals utilized in this study. The time required to perform each step is included in the respective steps.
1. Preparation of electrodes (Figure 1)
<ol>
	<li>Cut an appropriate length of tungsten wire (~20 mm) and burn off the insulation layer at one end to expose the tungsten wire (1.6-2 mm). Bend the other end of the wire into an &#34;L&#34; shape (see Figure 1A, ~1 min)</li>
	<li>Carefully place the &#34;L&#34; end of the tungsten wire onto the optical fiber (see Figure 1B, ~1 min). 
	NOTE: The tungsten wire should be 0.2-0.4 mm longer than the tip of optical fiber.</li>
	<li>Dip a drop of adhesive and apply it to the interface of the optical fiber and the tungsten wire (see Figure 1C, ~1 min). Due to the surface tension of the liquid, the tungsten wire will attach to the optical fiber.</li>
	<li>Apply the adhesive once and allow it to dry. Then, apply a second coat. This will further tighten the bond between the tungsten wire and the optical fiber (see Figure 1C, ~1 min). 
	NOTE: During the application process, care should be taken to prevent the adhesive from flowing to the tip of the optical fiber, as this may affect its transmittance.</li>
	<li>At the corner of the &#34;L&#34; shape, use adhesive to closely bond the base of the optical fiber and the tungsten wire together (~1 min). 
	NOTE: The increased contact area and the spherical solidification of the adhesive droplet ensure a firm bond between the optical fiber and the tungsten wire.</li>
	<li>Use soldering to establish a connection between the tungsten wire and the pin of the female connector (see Figure 1D, ~3 min for each) 
	NOTE: Since tungsten wire is not sensitive to heat, the success rate of soldering can be increased to nearly 100% by a metal sleeve fashioned by cutting the pin slot location to physically unite the two components. Subsequently, solder filling the sleeve ensures a stable connection between the inserted tungsten wire and the pin.</li>
	<li>According to the experimental design, solder the required number of optrodes (see Figure 1E, ~10 min) 
	NOTE: Three optrodes were utilized in this study.</li>
	<li>Cut the enameled wire to the appropriate length (~30 mm) and burn off the insulation on both ends. Wrap one end of the enameled wire around the base of the screw and solder it. Connect the other end of the enameled wire to the pins of the female connector via a soldering iron, serving as the EEG recording and ground connection for the skull electrode (see Figure 1F, ~5 min) 
	NOTE: Due to the flexibility of the enameled wire, it is convenient to connect the skull and the pins of the female connector. Soldering after winding the enameled wire around a screw is more reliable than surface-mount soldering.</li>
	<li>Apply hot-melt adhesive to the female connector&#39;s pin array to encapsulate the female connector and ensure the solder joints are fully enveloped. During the solidification of the hot-melt adhesive, shape the adhesive using a plastic board to minimize space occupancy during implantation (see Figure 1F, ~3 min) 
	NOTE: Due to the potential burn risks associated with using hot-melt glue, AB adhesive is considered a safer alternative.</li>
	<li>Next, use a light intensity meter and digital multimeter to measure luminousness and conductivity. Upon passing these tests, the setup is ready for use (see Figure 1G, ~2 min) 
	NOTE: When working with lasers, always wear protective eyewear suitable for the specific wavelength and power of the laser light. High-quality fibers are required to transmit at least 80% of the light from the end of the patch cord to the tip of the implantable fiber. Additionally, the resistance from the tip of the tungsten wire to the pin header slot was measured with an acceptable range of 1-5 &#937;.</li>
	<li>Before use, immerse the prepared electrodes in 75% alcohol for 15 min, then transfer them to sterile saline (~16 min). 
	NOTE: The total weight of the encapsulated electrode device (0.6701 &#177; 0.01123 g, n = 4) does not affect the activity of the mice. The electrodes are sterilized in advance using plasma sterilization.</li>
</ol>
2. Protocol for electrode implantation (Figure 2)
<ol>
	<li>Anesthetize the mice with isoflurane (2%-4%) and secure them in a stereotaxic apparatus (see Figure 2A, ~2 min). Use eye ointment on mouse eyes to prevent dryness while under anesthesia. Apply lidocaine to the skin around the perimeter of the cranial incision and a subcutaneous injection of carprofen at a dose of 5 mg/kg. 
	NOTE: Proper anesthetization is confirmed through the absence of the righting reflex and the loss of response to toe pinch. For prolonged surgical procedures, using an injectable&#160;anesthetic is more appropriate.</li>
	<li>After securing the animal in the stereotaxic apparatus, trim away the scalp along the edge of the skull and wipe the skull surface with 75% alcohol to expose the positions of the bregma and lambda (see Figure 2B, ~3 min). 
	NOTE: The pre-surgical procedure includes clipping the hair, disinfecting the surgical site, pre-sterilizing the instruments and surgical tools.</li>
	<li>Level the brain and drill holes (~0.8 mm) above the regions of interest (ROI) using a microdrill. Inject the relevant viruses with a syringe pump (see Figure 2C - F, ~20 min). 
	NOTE: Detailed information about the viruses and ROIs in this study is shown in the Table of Materials and Figure 3.</li>
	<li>Hold the optrodes with a holder and move their tip to the bregma point, setting this position as &#39;zero&#39;. Using the bregma point as the reference coordinate, move the optrodes to the target regions (see Figure 2G, ~3 min). Position the optrodes 0.3 mm above the virus injection site. 
	NOTE: In the vertical implantation strategy, the minimum&#160;distance between the two electrodes is approximately 2 mm, depending on the thickness of the fiber optic sleeve and the outer diameter of the ferrule. An angled insertion method can be used for simultaneous recording from closely located regions.</li>
	<li>After inserting the optic fibers into the corresponding brain areas, apply a small amount of tissue adhesive around them to cover the exposed tissue. Then, sequentially apply additional adhesive and a little dental cement. After the dental cement solidifies, release the holder carefully (see Figure 2H, ~5 min).</li>
	<li>Before each new optrode implantation, perform recalibration, which involves repeating step 2.4 (see Figure 2I). 
	NOTE: For each implantation, position the optrode at bregma as the &#39;zero&#39; point.</li>
	<li>After implanting all optrodes, drill the holes behind the lambda point for &#39;Ground&#39; and above the neocortex for &#39;EEG&#39; using a microdrill (~1 min). 
	NOTE: The size of the drilled hole (~1.1 mm) needs to match the diameter of the screw.</li>
	<li>Insert skull screws into the hole above the cortex and behind the lambda point (see Figure 2J, ~5 min). 
	NOTE: Skull screws are used for EEG recording and grounding, respectively.</li>
	<li>Organize the tungsten wires and enameled wires so they are positioned above the implantation area and not exposed outside the skull. Secure the position of the female connector using a holder (see Figure 2K, ~2 min).</li>
	<li>Fill the implantation site with dental cement to ensure a stable connection between the electrode device and the skull. Encapsulate the internal wires, leaving the female connector&#39;s insertion holes exposed for connection to the headstage (see Figure 2L, ~6 min).</li>
	<li>After dental cement hardening, apply iodine to the skin around the perimeter of the exposed scalp and then place the animal on a warming pad (~1 min). 
	NOTE: Ensure that the animal is not left unattended until it has regained sufficient consciousness to maintain sternal recumbency. Sterile conditions should be maintained throughout the survival surgery.</li>
	<li>Administer antibiotics and analgesics to support post-surgical recovery. 
	NOTE: Animals that have undergone surgery are not reintroduced to the company of other animals until they have fully recovered.Animals were administered intraperitoneal injections of ceftriaxone sodium (180 mg/kg),&#160;and subcutaneous injections of carprofen (5 mg/kg),&#160;and lidocaine gel was applied to the surgical wounds daily for 3 days. The animals are required to survive for a designated period to complete the behavioral testing. Subsequently, they will be euthanized using 4%-5% isoflurane, followed by perfusion and tissue sectioning to assess viral infection and the sites of electrode implantation.</li>
</ol>
3. Protocol for recording in vivo brain
<ol>
	<li>Before conducting signal recording and behavioral testing (open field test)15, ensure that the mouse undergoes a 3-day period of gentle handling to acclimatize them to the environment. 
	NOTE: In this study, the mice were placed in an open field (42 cm &#215; 42 cm) to move around freely, allowing us to observe their seizure behavior.</li>
	<li>After anesthetizing the mice with isoflurane (2%-4%), connect the optical fibers sequentially in the order of the recording site before inserting the headstage.</li>
	<li>Set the 635 nm light pulse with a 10% duty cycle at 20 Hz. Set up the light stimulation for 10 s (200 pulses) during recording. Set the intensity at the optical fiber tips at 3 mW.</li>
	<li>Use a fiber photometry system to record Gcamp6m signals behavioral testing period at a sampling rate of 30 Hz by 470 nm LED channel.</li>
	<li>Simultaneously record calcium signals, LFPs, and EEG under optogenetic stimulation, along with corresponding behavioral performances (see Figure 3).</li>
	<li>Employ transistor-transistor logic (TTL) signal synchronization tagging to ensure temporal consistency in recorded data.</li>
</ol>
4. Data processing
<ol>
	<li>Load photometry data to MATLAB software. Then, use custom coding (Supplementary File 1) to detrend the calcium signal.</li>
	<li>Define the changes in fluorescence intensity with equation (1): 
	&#916;F/F = (Ft&#160;-&#160;F0)/ F0&#160; &#160; (1) 
	Where Ft represents the real-time calcium signal fluorescence values, and F0 denotes the averaged baseline fluorescence intensity before the event.</li>
	<li>Import the electrophysiological data, sampled rate of 1000 Hz, into MATLAB (Supplementary File 2).</li>
	<li>Coordinate the display of calcium signaling and electrophysiological data in alignment with synchronized markers.</li>
</ol>

Flexible Multi-Region Neural Recording Device for Epilepsy Research

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The authors declare no competing financial interests to disclose.

Here, we employed a self-made optrode device for in vivo neural signal recording across multiple regions. The feasibility of this system for simultaneous optogenetic stimulation, calcium signal recording, and electrophysiological recording has been validated. The electrode preparation method described herein is efficient and cost-effective. According to the experimental design, we could record signals from relevant brain regions. The strategic arrangement of optrodes allows for the potential extraction of abunda...

Epilepsy is a common neurological condition characterized by recurrent seizures, which manifest as convulsions, sensory disturbances, and loss of consciousness1. The pathophysiological mechanisms underlying epilepsy are complex and involve multiple interconnected brain regions2,3. Recent advances in neuroimaging have shed light on the large-scale networks involved in epilepsy4,5. However, understanding of the intricate circuitry and network mechanisms underlying the generation and propagation of epilepsy remains limited, partly due to the insufficient application of multi-region neural recording techniques6. Therefore, developing a flexible, integrated method capable of simultaneously monitoring neural activity across disparate brain regions is imperative.
Electrophysiological recordings are conducted to capture seizures and determine the presence of epilepsy7. Except for recording electrophysiological activity, there is a growing emphasis on the precise calcium activity of specific&#160;neural populations in epilepsy studies8,9. Advancements in calcium indicator synthesis and various probe designs have prompted researchers to adopt fiber photometry for capturing changes in neuronal activity and neural substances within the brain10,11. The two independent methods of detecting neuronal activity, namely electrophysiology and fiber photometry recording, complement each other, facilitating a more comprehensive understanding of dynamic neuronal processes.
Additionally, synchronous recording and modulating neural activity are pivotal for gaining insights into the brain functioning at both network and cellular levels. This approach enables researchers to observe and manipulate the brain&#39;s complex processes in real-time. Optogenetics has emerged as an indispensable tool for probing neural signaling, owing to its distinctive ability for selective stimulation or inhibition12. Despite widespread applications of multi-site electrophysiological recording in neuroscience13, the integration of multi-region electrophysiological recording with fiber photometry and optogenetic manipulation remains limited. More importantly, current methods for fabricating and implanting multi-region electrodes lack flexibility14. These limitations hinder our capacity to dissect specific circuit functions and interactions across multiple regions. Here, we present a cost-effective and convenient multi-region in vivo recording approach that sheds light on neuronal processes across regions in kindling-induced seizures and other neuropsychiatric disorders.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>8-32 adapter</td>
			<td>Plexon</td>
			<td>Custom ordered</td>
			<td>Connect the female connector and headstage</td>
		</tr>
		<tr>
			<td>AAV-CaMKII&#945;-ChrimsonR-mcherry</td>
			<td>Taitool Bioscience</td>
			<td>S0371-9</td>
			<td>4 x 1012 VG/mL&#160;</td>
		</tr>
		<tr>
			<td>AAV-hsyn-Gcamp6m</td>
			<td>Taitool Bioscience</td>
			<td>S0471-9</td>
			<td>4 x 1012 VG/mL&#160;</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma</td>
			<td>236276</td>
			<td>Titered 1:500</td>
		</tr>
		<tr>
			<td>Dental Cement</td>
			<td>New Century Dental</td>
			<td>430205</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrophysiological recordings system</td>
			<td>Plexon</td>
			<td>Omniplex</td>
			<td></td>
		</tr>
		<tr>
			<td>Enameled wire</td>
			<td>N/A</td>
			<td>Custom ordered</td>
			<td>Diameter = 0.2 mm</td>
		</tr>
		<tr>
			<td>Female connector</td>
			<td>N/A</td>
			<td>Custom ordered</td>
			<td>1.25 mm pitch</td>
		</tr>
		<tr>
			<td>Glue</td>
			<td>Loctite</td>
			<td>45282</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser</td>
			<td>Changchun New Industries</td>
			<td>BH81563</td>
			<td>635 nm&#160;</td>
		</tr>
		<tr>
			<td>MATLAB</td>
			<td>MathWorks</td>
			<td>R2021b</td>
			<td></td>
		</tr>
		<tr>
			<td>Microdrill</td>
			<td>RWD</td>
			<td>78001</td>
			<td></td>
		</tr>
		<tr>
			<td>Multichannel fiber photometry</td>
			<td>ThinkerTech</td>
			<td>FPS-SS-MC-LED</td>
			<td></td>
		</tr>
		<tr>
			<td>Optical fiber</td>
			<td>Xi&#39;an Bogao</td>
			<td>L-200UM</td>
			<td>Select the appropriate fiber length based on the depth of the targeted brain regions.</td>
		</tr>
		<tr>
			<td>PFA-Coated Tungsten wire</td>
			<td>A-M System</td>
			<td>795500</td>
			<td>Bare 0.002&#34;; Coated 0.0040&#34;</td>
		</tr>
		<tr>
			<td>Power meter</td>
			<td>Thorlabs</td>
			<td>PM100D</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic Fxrame</td>
			<td>RWD</td>
			<td>68807</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue adhesive</td>
			<td>3M</td>
			<td>1469SB</td>
			<td></td>
		</tr>
	</tbody>
</table>

an integrated method for crafting flexible and convenient electrophysiological optrodes for multi-region in vivo recording

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

We combined optogenetics with multiregional electrophysiological recording and calcium imaging to observe neuronal activity across various brain regions during optogenetic seizures. For this purpose, an adeno-associated virus (AAV) expressing ChrimsonR under the control of the CaMKII&#945; promoter (AAV-CaMKII&#945;-ChrimsonR-mcherry)16 was injected into a classical epileptogenic site, the piriform cortex (ROI 1)17, in rodents. Additionally, AAV-hsyn-Gcamp6m<sup class="xref...

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

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

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

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

an-integrated-method-for-crafting-flexible-convenient

Research

JoVE Journal

Neuroscience

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

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

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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 ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

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

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

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

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

A Method for Systematic Electrochemical and Electrophysiological Evaluation of Neural Recording Electrodes

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant characteristics.&#160;This precludes a rational selection of a particular implant as optimal for a particular application and the development of new materials based on the most critical performance parameters.&#160;This article develops a protocol for in vitro and in vivo testing of neural recording electrodes.&#160;Recommended parameters for electrochemical and electrophysiological testing are documented with the key steps and potential issues discussed.&#160;This method eliminates or reduces the impact of many systematic errors present in simpler in vivo testing paradigms, especially variations in electrode/neuron distance and between animal models.&#160;The result is a strong correlation between the critical in vitro and in vivo responses, such as impedance and signal-to-noise ratio.&#160;This protocol can easily be adapted to test other electrode materials and designs.&#160;The in vitro techniques can be expanded to any other nondestructive method to determine further important performance indicators.&#160;The principles used for the surgical approach in the auditory pathway can also be modified to other neural regions or tissue.

Different electrode coatings affect neural recording performance through changes to electrochemical, chemical and mechanical properties. Comparison of electrodes in vitro is relatively simple, however comparison of in vivo response is typically complicated by variations in electrode/neuron distance and between animals. This article provides a robust method to compare neural recording electrodes.

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of primary afferent fibers mediated by GABAergic axo-axonal synapses (primary afferent depolarization). The strength of primary afferent depolarization can be measured by recording of volume-conducted potentials at the dorsal root (dorsal root potentials, DRP). Pathological changes of presynaptic inhibition are crucial in the abnormal central processing of certain pain conditions and in some disorders of motor hyperexcitability. Here, we describe a method of recording DRP in vivo in mice. The preparation of spinal cord dorsal roots in the anesthetized animal and the recording procedure using suction electrodes are explained. This method allows measuring GABAergic DRP and thereby estimating spinal presynaptic inhibition in the living mouse. In combination with transgenic mouse models, DRP recording may serve as a powerful tool to investigate disease-associated spinal pathophysiology. In vivo recording has several advantages compared to ex vivo isolated spinal cord preparations, e.g. the possibility of simultaneous recording or manipulation of supraspinal networks and induction of DRP by stimulation of peripheral nerves.

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

GABAergic presynaptic inhibition is a powerful inhibitory mechanism in the spinal cord important for motor and sensory signal integration in spinal cord networks. Underlying primary afferent depolarization can be measured by recording of dorsal root potentials (DRP). Here we demonstrate a method of in vivo recording of DRP in mice.

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of ...

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.