Name: recording network activity in spinal nociceptive circuits using microelectrode arrays
Uploaded: 2022-02-09T12:30:00
Description: Watch this Scientific Journal Video about Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays at JoVE.com

This work was funded by the National Health and Medical Research Council (NHMRC) of Australia (grants 631000, 1043933, 1144638, and 1184974 to B.A.G. and R.J.C.) and the Hunter Medical Research Institute (grant to B.A.G. and R.J.C.).

Studies were carried out on male and female c57Bl/6 mice aged 3-12 months. All experimental procedures were performed in accordance with the University of Newcastle&#39;s Animal Care and Ethics Committee (protocols A-2013-312, and A-2020-002).
1. In vitro electrophysiology
<ol>
	<li>Preparation of solutions for spinal cord slice preparation and recording
	<ol>
		<li>Artificial cerebrospinal fluid 
		&#8203;NOTE: Artificial cerebrospinal fluid (aCSF) is used in an interface incu...

<ol>
	<li>Smith, K. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calretinin+positive+neurons+form+an+excitatory+amplifier+network+in+the+spinal+cord+dorsal+horn.">Calretinin positive neurons form an excitatory amplifier network in the spinal cord dorsal horn.</a> eLife. 8, 49190 (2019).</li><li>Smith, K. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+heterogeneity+of+calretinin-expressing+neurons+in+the+mouse+superficial+dorsal+horn:+implications+for+spinal+pain+processing.">Functional heterogeneity of calretinin-expressing neurons in the mouse superficial dorsal horn: implications for spinal pain processing.</a> The Journal of physiology. 593 (19), 4319-4339 (2015).</li><li>Boyle, K. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Defining+a+spinal+microcircuit+that+gates+myelinated+afferent+input:+Implications+for+tactile+allodynia.">Defining a spinal microcircuit that gates myelinated afferent input: Implications for tactile allodynia.</a> Cell Reports. 28 (2), 526-540 (2019).</li><li>Browne, T. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transgenic+cross-referencing+of+inhibitory+and+excitatory+interneuron+populations+to+dissect+neuronal+heterogeneity+in+the+dorsal+horn.">Transgenic cross-referencing of inhibitory and excitatory interneuron populations to dissect neuronal heterogeneity in the dorsal horn.</a> Frontiers in Molecular Neuroscience. 13, 32 (2020).</li><li>Graham, B. A., Hughes, D. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rewards,+perils+and+pitfalls+of+untangling+spinal+pain+circuits.">Rewards, perils and pitfalls of untangling spinal pain circuits.</a> Current Opinion in Physiology. 11, 35-41 (2019).</li><li>Todd, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+circuitry+for+pain+processing+in+the+dorsal+horn.">Neuronal circuitry for pain processing in the dorsal horn.</a> Nature Reviews Neuroscience. 11 (12), 823-836 (2010).</li><li>Hughes, D. I., Todd, A. 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Despite the importance of the spinal DH in nociceptive signaling, processing, and the resulting behavioral and emotional responses that characterize pain, the circuits within this region remain poorly understood. A key challenge in investigating this issue has been the diversity of neuron populations that comprise these circuits6,31,32. Recent advances in transgenic technologies, led by optogenetics and chemogenetics, are beginn...

The roles of specific types of inhibitory and excitatory interneurons within the spinal cord DH are being uncovered at a rapid rate1,2,3,4. Together, interneurons make up over 95% of the neurons in the DH and are involved in sensory processing, including nociception. Furthermore, these interneuron circuits are important for determining whether peripheral signals ascend the neuroaxis to reach the brain and contribute to the perception of pain5,6,7. To date, most studies have investigated the role of DH neurons at either the single-cell or whole-organism level of analysis using combinations of in vitro intracellular electrophysiology, neuroanatomical labeling, and in vivo behavioral analysis1,3,8,9,10,11,12,13,14. These approaches have significantly advanced the understanding of the role of specific neuron populations in pain processing. However, a gap remains in understanding how specific cell types and small macro-circuits influence large populations of neurons at a microcircuit level to subsequently shape the output of the DH, behavioral responses, and the pain experience.
One technology that can investigate macro-circuit or multicellular-level function is the microelectrode array (MEA)15,16. MEAs have been used to investigate nervous system function for several decades17,18. In the brain, they have facilitated the study of neuronal development, synaptic plasticity, pharmacological screening, and toxicity testing17,18. They can be used for both in vitro and in vivo applications, depending on the type of MEA. Furthermore, the development of MEAs has evolved rapidly, with different electrode numbers and configurations now available19. A key advantage of MEAs is their capacity to simultaneously assess electrical activity in many neurons with high spatial and temporal accuracy via multiple electrodes15,16. This provides a broader readout of how neurons interact in circuits and networks, under control conditions and in the presence of locally applied compounds.
One challenge of in vitro DH preparations is that ongoing activity levels are typically low. Here, this challenge is addressed in spinal cord DH circuits using the voltage-gated K+ channel blocker, 4-aminopryidine (4-AP), to chemically stimulate DH circuits. This drug has previously been used to establish rhythmic synchronous electrical activity in the DH of acute spinal cord slices and under acute in vivo conditions20,21,22,23,24. These experiments have used single-cell patch and extracellular recording or calcium imaging to characterize 4-AP-induced activity20,21,22,23,24,25. Together, this work has demonstrated the requirement of excitatory and inhibitory synaptic transmission and electrical synapses for rhythmic 4-AP-induced activity. Thus, the 4-AP response has been viewed as an approach that unmasks native polysynaptic DH circuits with biological relevance rather than as a drug-induced epiphenomenon. Furthermore, 4-AP-induced activity exhibits a similar response profile to analgesic and antiepileptic drugs as neuropathic pain conditions and has been used to propose novel spinally-based analgesic drug targets such as connexins20,21,22.
Here, a preparation that combines MEAs and chemical activation of the spinal DH with 4-AP to study this nociceptive circuitry at the macro-circuit, or network level of analysis, is described. This approach provides a stable and reproducible platform for investigating nociceptive circuits under naive and neuropathic &#39;pain-like&#39; conditions. This preparation is also readily applicable to test the circuit-level action of known analgesics and to screen novel analgesics in the hyperactive spinal cord.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>4-aminopyridine</td>
			<td>Sigma-Aldrich</td>
			<td>275875-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>100% ethanol</td>
			<td>Thermo Fisher</td>
			<td>AJA214-2.5LPL</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2 1M</td>
			<td>Banksia Scientific</td>
			<td>0430/1L</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbonox (Carbogen - 95% O2, 5% CO2)</td>
			<td>Coregas</td>
			<td>219122</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved long handle spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15015-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom made air interface incubation chamber</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Foetal bovine serum</td>
			<td>Thermo Fisher</td>
			<td>10091130</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps Dumont #5</td>
			<td>Fine Science Tools</td>
			<td>11251-30</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Thermo Fisher</td>
			<td>AJA783-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Horse serum</td>
			<td>Thermo Fisher</td>
			<td>16050130</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted microscope</td>
			<td>Zeiss</td>
			<td>Axiovert10</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Thermo Fisher</td>
			<td>AJA383-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>Ceva</td>
			<td>KETALAB04</td>
			<td></td>
		</tr>
		<tr>
			<td>Large surgical scissors</td>
			<td>Fine Science Tools</td>
			<td>14007-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Loctite 454 Instant Adhesive</td>
			<td>Bolts and Industrial Supplies</td>
			<td>L4543G</td>
			<td></td>
		</tr>
		<tr>
			<td>MATLAB</td>
			<td>MathWorks</td>
			<td></td>
			<td>R2018b</td>
		</tr>
		<tr>
			<td>MEAs, 3-Dimensional</td>
			<td>Multichannel Systems</td>
			<td>60-3DMEA100/12/40iR-Ti, 60-3DMEA200/12/50iR-Ti</td>
			<td>60 titanium nitride (TiN) electrodes with 1 internal reference electrode, organised in an 8x8 square grid. Electrodes are 12 &#181;m in diameter, 40 &#181;m (100/12/40) or 50 &#181;m (200/12/50) high and equidistantly spaced 100 &#181;m (100/12/40) or 200 &#181;m (200/12/50) apart.</td>
		</tr>
		<tr>
			<td>MEA headstage</td>
			<td>Multichannel Systems</td>
			<td>MEA2100-HS60</td>
			<td></td>
		</tr>
		<tr>
			<td>MEA interface board</td>
			<td>Multichannel Systems</td>
			<td>MCS-IFB 3.0 Multiboot</td>
			<td></td>
		</tr>
		<tr>
			<td>MEA net</td>
			<td>Multichannel Systems</td>
			<td>ALA HSG-MEA-5BD</td>
			<td></td>
		</tr>
		<tr>
			<td>MEA perfusion system</td>
			<td>Multichannel Systems</td>
			<td>PPS2</td>
			<td></td>
		</tr>
		<tr>
			<td>MEAs, Planar</td>
			<td>Multichannel Systems</td>
			<td>60MEA200/30iR-Ti, 60MEA500/30iR-Ti</td>
			<td>60 titanium nitride (TiN) electrodes with 1 internal reference electrode, organised in either a 8x8 square grid (200/30) or a 6x10 rectangular grid (500/30). Electrodes are 30 &#181;m in diameter and equidistantly spaced 200 &#181;m (200/30) or 500 &#181;m (500/30) apart.</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Thermo Fisher</td>
			<td>AJA296-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope camera</td>
			<td>Motic</td>
			<td>Moticam X Wi-Fi</td>
			<td></td>
		</tr>
		<tr>
			<td>Multi Channel Analyser software</td>
			<td>Multichannel Systems</td>
			<td></td>
			<td>V 2.17.4</td>
		</tr>
		<tr>
			<td>Multi Channel Experimenter software</td>
			<td>Multichannel Systems</td>
			<td></td>
			<td>V 2.17.4</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Thermo Fisher</td>
			<td>AJA465-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Thermo Fisher</td>
			<td>AJA475-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Thermo Fisher</td>
			<td>ACR207805000</td>
			<td></td>
		</tr>
		<tr>
			<td>Rongeurs</td>
			<td>Fine Science Tools</td>
			<td>16021-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Small spring scissors</td>
			<td>Fine Science Tools</td>
			<td>91500-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Small surgical scissors</td>
			<td>Fine Science Tools</td>
			<td>14060-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Thermo Fisher</td>
			<td>AJA530-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Superglue</td>
			<td></td>
			<td></td>
			<td>cyanoacrylate adhesive</td>
		</tr>
		<tr>
			<td>Tetrodotoxin</td>
			<td>Abcam</td>
			<td>AB120055</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibration isolation table</td>
			<td>Newport</td>
			<td>VH3048W-OPT</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibrating microtome</td>
			<td>Leica</td>
			<td>VT1200 S</td>
			<td></td>
		</tr>
	</tbody>
</table>

recording network activity in spinal nociceptive circuits using microelectrode arrays

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an increasingly detailed view of the circuits underpinning spinal pain processing. However, the effects of these connections for broader network activity in the DH remain less well understood because most studies focus on the activity of single neurons and small microcircuits. Alternatively, the use of microelectrode arrays (MEAs), which can monitor electrical activity across many cells, provides high spatial and temporal resolution of neural activity. Here, the use of MEAs with mouse spinal cord slices to study DH activity induced by chemically stimulating DH circuits with 4-aminopyridine (4-AP) is described. The resulting rhythmic activity is restricted to the superficial DH, stable over time, blocked by tetrodotoxin, and can be investigated in different slice orientations. Together, this preparation provides a platform to investigate DH circuit activity in tissue from na&#239;ve animals, animal models of chronic pain, and mice with genetically altered nociceptive function. Furthermore, MEA recordings in 4-AP-stimulated spinal cord slices can be used as a rapid screening tool to assess the capacity of novel antinociceptive compounds to disrupt activity in the spinal cord DH.

Model of network activity in the spinal cord dorsal horn 
Application of 4-AP reliably induces synchronous rhythmic activity in the spinal cord DH. Such activity presents as increased EAPs and LFPs. The later signal is a low-frequency waveform, which has previously been described in MEA recordings30. Changes in EAP and/or LFP activity following drug application reflect altered neural activity. Examples of EAPs and LFPs are shown in Figure 3B and ...

Watch this Scientific Journal Video about Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays at JoVE.com

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

The combined use of microelectrode array technology and 4-aminopyridine-induced chemical stimulation for investigating network-level nociceptive activity in the spinal cord dorsal horn is outlined.

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an ...

The MEA 4-AP spinal slice preparation described to you provides a platform to study the connectivity of dorsal horn circuits and how these networks interact during spinal sensory processing. The methodology presented enables dorsal horn circuit activity to be investigated at a macroscopic regional level of resolution. This preparation can be used as a rapid screening tool to assess the ability of compounds to disrupt signaling in spinal sensory circuits.This method helps bridge the gap in our understanding of how the activity of specific cell types and small micro circuits influence large populations of neurons to subsequently shape the output of the dorsal horn behavior responses and ultimately the pain experience. To begin, fill the MEA well with horse serum for 30 minutes. After removing the serum, thoroughly rinse the MEA five times with distilled water until the distilled water becomes non-foamy.Then, fill the well with artificial CSF. After decapitating the anesthetized mouse, remove the skin over the abdominal region by making a small cut in the skin at the level of the hips. Then, pull the skin on either side of the cut rostrally until all the skin is removed from the top of the rib cage to the top of the pelvis, both ventrally and dorsally.After placing the body on ice, use a ventral approach to expose the vertebral column by removing the viscera and cutting through the ribs laterally to the sternum. Remove the ventral rib cage, both scapulae, and the lower limbs and pelvis. Transfer the vertebral column and rib preparation to a dissecting bath containing ice cold sucrose artificial CSF.Pin all four corners of the preparation by placing pins through the lower back muscles and the attached upper ribs. Remove all muscle and connective tissue overlying the ventral surface of the vertebrae with the rongeurs and identify the vertebral region over the lumbo-sacral enlargement, which lies approximately beneath the T12-L2 vertebral bodies. Remove a vertebral body caudal to the lumbo-sacral enlargement region to access the spinal cord sitting on the vertebral canal.Using curved spring scissors, cut through the vertebral pedicles bilaterally while lifting and pulling the vertebral body rostrally to separate the ventral and dorsal aspects of the vertebrae and expose the spinal cord. Once the vertebral bodies are removed to reveal the lumbo-sacral enlargement, carefully clear the remaining roots that anchor the spinal cord with spring scissors until the cord floats free. Isolate the spinal cord with rostral and caudal cuts above and below the lumbo-sacral enlargement to make the target region of the cord float free.For transverse slices, lift and place the lumbo-sacral segment by an attached route on a precut polystyrene block with a shallow channel cut in the center. Then, use cyanoacrylate adhesive to attach the block and cord to the sectioning platform and place it in the cutting bath containing ice cold sucrose artificial CSF slurry. Once the 300 micrometers thick slices are obtained, transfer the slices to an air interface incubation chamber containing oxygenated artificial CSF and allow the slices to equilibrate for one hour at room temperature.To begin recording dorsal horn activity, transfer the slice from the incubator to the MEA well using a large tip pasteur pipette filled with artificial CSF and add additional artificial CSF. Use a fine short hair paint brush to position the slice over the 60 electrode recording array. Then, place a weighted neck over the tissue to hold it in place and promote good contact with MEA electrodes.Place the MEA in the recording head stage. Check the position of the tissue over the electrodes using an inverted microscope to confirm that as many electrodes as possible are under the superficial DH.Ensure that at least two to six electrodes do not contact the slice. After connecting the camera to the device, take a reference image of the slice relative to the MEA for use during the analysis.Then, press start DAQ in the recording software and confirm that all electrodes receive a clear signal. Next, attach the perfusion inlet and outlet lines to the MEA well filled with artificial CSF and turn the perfusion system on. Check the flow rate and ensure that the outflow is sufficient to prevent overflow of the superfusate.After equilibrating the tissue for five minutes, record the raw baseline data for five minutes. Move the perfusion inlet line from artificial CSF to a 4-aminopyridine solution and wait for 12 minutes for the 4-aminopyridine induced rhythmic activity to reach steady state. Then, record five minutes of 4-aminopyridine induced activity and be prepared for the subsequent recordings to test the drugs or check the stability of 4-aminopyridine.Following each recording session, rinse the lines with artificial CSF. After removing the MEA from the head stage and removing the net and the tissue from the MEA well, rinse the MEA and net with artificial CSF and repeat the whole process with a new slice. Open the analysis software and load the pre-made analysis layout.Open the file of interest and deselect the reference electrode and any electrodes deemed to be excessively noisy. Set the time window for analysis. Then, move to the cross channel filter tab.Select the complex reference and reference electrodes based on the image taken and notes made during the experiment and press Explore before continuing. Move to the EAP filter tab and apply a second order high-pass Butterworth filter to remove LFP activity. Move to the LFP filter tab and apply a second order band pass Butterworth filter to remove EAP activity.In the EAP detector tab, ensure to select auto threshold, tick the rising and falling edge boxes, and set the dead time to three milliseconds. To set positive and negative thresholds, inspect the data by returning to the raw data analyzer screen, moving the time marker, returning to the EAP detector tab, and pressing explore. Repeat the process until the set detection threshold captures EAPs without capturing noise or non-physiological activity.In the LFP detector tab, ensure to select the manual threshold, tick the rising and falling edge boxes, and set the dead time to three milliseconds. Set threshold for one electrode and, once satisfied, select apply to all and then press start analysis. The bath application of the voltage gated sodium channel antagonist tetrodotoxin abolished both EAP and LFP activity, confirming the spike dependency of these signals.The stability of 4-aminopyridine induced activity parameters was characterized for EAP or LFP. All activity characteristics plus the coincidence of activity for LFPs were stable based on the similarity of 4-aminopyridine induced activity at 12 minutes after 4-aminopyridine application and 15 minutes later. The feature's EAPs or LFPs were characterized by an increase in the number of coincident events detected across multiple electrodes.The number of linked adjacent electrodes, and the strength of linkages between adjacent electrodes and LFPs following 4-aminopyridine stimulation and then relative stability. The slice orientation is an important consideration for in vitro preparations. 4-aminopyridine stimulation induced a similar rhythmic activity in the SDH regardless of slice orientation.Finally, to shed further light on the relevance of 4-aminopyridine activation of the DH networks, an elevated potassium artificial CSF solution was bath applied and shown to evoke a similar DH response to 4-aminopyridine stimulation. Key steps in the protocol are in the tissue preparation and placement of sections, ensuring the tissue is healthy through careful, but timely, dissection, as well as delicate optimal positioning of the tissue on the MEA will aid in obtaining quality results.

recording-network-activity-spinal-nociceptive-circuits-using

Research

JoVE Journal

Neuroscience

Scalable Fluidic Injector Arrays for Viral Targeting of Intact 3-D Brain Circuits

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted in neurological and psychiatric disorders--would be greatly facilitated by a technology for rapidly targeting genes to complex 3-dimensional neural circuits, enabling fast creation of "circuit-level transgenics." We have recently developed methods in which viruses encoding for light-sensitive proteins can sensitize specific cell types to millisecond-timescale activation and silencing in the intact brain. We here present the design and implementation of an injector array capable of delivering viruses (or other fluids) to dozens of defined points within the 3-dimensional structure of the brain (Figure. 1A, 1B). The injector array comprises one or more displacement pumps that each drive a set of syringes, each of which feeds into a polyimide/fused-silica capillary via a high-pressure-tolerant connector. The capillaries are sized, and then inserted into, desired locations specified by custom-milling a stereotactic positioning board, thus allowing viruses or other reagents to be delivered to the desired set of brain regions. To use the device, the surgeon first fills the fluidic subsystem entirely with oil, backfills the capillaries with the virus, inserts the device into the brain, and infuses reagents slowly (&lt;0.1 microliters/min). The parallel nature of the injector array facilitates rapid, accurate, and robust labeling of entire neural circuits with viral payloads such as optical sensitizers to enable light-activation and silencing of defined brain circuits. Along with other technologies, such as optical fiber arrays for light delivery to desired sets of brain regions, we hope to create a toolbox that enables the systematic probing of causal neural functions in the intact brain. This technology may not only open up such systematic approaches to circuit-focused neuroscience in mammals, and facilitate labeling of brain regions in large animals such as non-human primates, but may also open up a clinical translational path for cell-specific optical control prosthetics, whose precision may enable improved treatment of intractable brain disorders. Finally, such devices as described here may facilitate precisely-timed fluidic delivery of other payloads, such as stem cells and pharmacological agents, to 3-dimensional structures, in an easily user-customizable fashion.

Controlling and analyzing neural circuits in vivo would be facilitated by a technology for delivery of viruses and other reagents to desired 3-dimensional sets of brain regions. We demonstrate customized fluidic injector array fabrication, and delivery of virally-encoded optical sensitizers, enabling optical manipulation of complex brain circuits.

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted ...

Examining Local Network Processing using Multi-contact Laminar Electrode Recording

Cortical layers are ubiquitous structures throughout neocortex1-4 that consist of highly recurrent local networks. In recent years, significant progress has been made in our understanding of differences in response properties of neurons in different cortical layers5-8, yet there is still a great deal left to learn about whether and how neuronal populations encode information in a laminar-specific manner.
Existing multi-electrode array techniques, although informative for measuring responses across many millimeters of cortical space along the cortical surface, are unsuitable to approach the issue of laminar cortical circuits. Here, we present our method for setting up and recording individual neurons and local field potentials (LFPs) across cortical layers of primary visual cortex (V1) utilizing multi-contact laminar electrodes (Figure 1; Plextrode U-Probe, Plexon Inc).
The methods included are recording device construction, identification of cortical layers, and identification of receptive fields of individual neurons. To identify cortical layers, we measure the evoked response potentials (ERPs) of the LFP time-series using full-field flashed stimuli. We then perform current-source density (CSD) analysis to identify the polarity inversion accompanied by the sink-source configuration at the base of layer 4 (the sink is inside layer 4, subsequently referred to as granular layer9-12). Current-source density is useful because it provides an index of the location, direction, and density of transmembrane current flow, allowing us to accurately position electrodes to record from all layers in a single penetration6, 11, 12.

A fundamental issue in our understanding of cortical circuitry is how networks in different cortical layers encode sensory information. Here we describe electrophysiological techniques utilizing multi-contact laminar electrodes to record single-units and local field potentials and present analyses to identify cortical layers.

Cortical layers are ubiquitous structures throughout neocortex1-4 that consist of highly recurrent local networks. In recent years, significant progress ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

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 ...

Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to stimuli remain to be fully understood. To overcome the difficulty of accessing the brainstem in the whole animal, isolation of the brainstem and part of the spinal cord is performed. This preparation is maintained in artificial cerebro-spinal fluid where gases, concentrations, and temperature are controlled and monitored. The output signal from the respiratory network is recorded by a suction electrode placed on the fourth ventral root. In this manner, stimuli can be directly applied onto the brainstem, and the effect can be recorded directly. The signal recorded is linked to the inspiratory signal sent to the diaphragm via the phrenic nerve, and can be described as bursts (around 8 bursts per minute). Analysis of these bursts (frequency, amplitude, length, and area under the curve) allows precise characterization of the stimulus effect on the respiratory network. The main limitation of this method is the viability of the preparation beyond the early post-natal stages. Thus, this method greatly focuses on the study of the whole network without the peripheral inputs in the newborn rat.

The central respiratory drive is located in the brainstem. Spontaneous respiratory motor output from an isolated brainstem-spinal cord is recorded by placing an electrode on the fourth ventral root. This experimental approach is valuable for pharmacological investigations or the assessment of respiratory challenges and genetic manipulations on rhythmic motor behavior.

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) is a promising approach when pharmacological treatment fails or neurosurgery is not recommended. Acute brain slices coupled to microelectrode arrays (MEAs) represent a valuable tool to study neuronal network interactions and their modulation by electrical stimulation. As compared to conventional extracellular recording techniques, they provide the added advantages of a greater number of observation points and a known inter-electrode distance, which allow studying the propagation path and speed of electrophysiological signals. However, tissue oxygenation may be greatly impaired during MEA recording, requiring a high perfusion rate, which comes at the cost of decreased signal-to-noise ratio and higher oscillations in the experimental temperature. Electrical stimulation further stresses the brain tissue, making it difficult to pursue prolonged recording/stimulation epochs. Moreover, electrical modulation of brain slice activity needs to target specific structures/pathways within the brain slice, requiring that electrode mapping be easily and quickly performed live during the experiment. Here, we illustrate how to perform the recording and electrical modulation of 4-aminopyridine (4AP)-induced epileptiform activity in rodent brain slices using planar MEAs. We show that the brain tissue obtained from mice outperforms rat brain tissue and is thus better suited for MEA experiments. This protocol guarantees the generation and maintenance of a stable epileptiform pattern that faithfully reproduces the electrophysiological features observed with conventional field potential recording, persists for several hours, and outlasts sustained electrical stimulation for prolonged epochs. Tissue viability throughout the experiment is achieved thanks to the use of a small-volume custom recording chamber allowing for laminar flow and quick solution exchange even at low (1 mL/min) perfusion rates. Quick MEA mapping for real-time monitoring and selection of stimulating electrodes is performed by a custom graphic user interface (GUI).

We illustrate how to perform recording and electrical modulation of 4-aminopyridine-induced epileptiform activity in rodent brain slices using microelectrode arrays. A custom recording chamber maintains tissue viability throughout prolonged experimental sessions. Live electrode mapping and selection of stimulating pairs are performed by a custom graphical user interface.

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) ...

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset (Callithrix jacchus)

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the neurosciences. Phylogenetically, these animals are much closer to humans than rodents. They also display complex behaviors, including a wide range of vocalizations and social interactions. Here, an effective stereotaxic neurosurgical procedure for implantation of recording electrode arrays in the common marmoset is described. This protocol also details the pre- and postoperative steps of animal care that are required to successfully perform such a surgery. Finally, this protocol shows an example of local field potential and spike activity recordings in a freely behaving marmoset 1 week after the surgical procedure. Overall, this method provides an opportunity to study the brain function in awake and freely behaving marmosets. The same protocol can be readily used by researchers working with other small primates. In addition, it can be easily modified to allow other studies requiring implants, such as stimulating electrodes, microinjections, implantation of optrodes or guide cannulas, or ablation of discrete tissue regions.

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset Callithrix jacchus

This work presents a protocol to perform a stereotaxic, neurosurgical implantation of&#160;microelectrode arrays in the common marmoset. This method specifically enables electrophysiological recordings in freely behaving animals but can be easily adapted to any other similar neurosurgical intervention in this species (e.g., cannula for drug administration or electrodes for brain stimulation).

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the ...

Multi-Fiber Photometry to Record Neural Activity in Freely-Moving Animals

Recording the activity of a group of neurons in a freely-moving animal is a challenging undertaking. Moreover, as the brain is dissected into smaller and smaller functional subgroups, it becomes paramount to record from projections and/or genetically-defined subpopulations of neurons. Fiber photometry is an accessible and powerful approach that can overcome these challenges. By combining optical and genetic methodologies, neural activity can be measured in deep brain structures by expressing genetically-encoded calcium indicators, which translate neural activity into an optical signal that can be easily measured. The current protocol details the components of a multi-fiber photometry system, how to access deep brain structures to deliver and collect light, a method to account for motion artifacts, and how to process and analyze fluorescent signals. The protocol details experimental considerations when performing single and dual color imaging, from either single or multiple implanted optic fibers.

This protocol details how to implement and perform multi-fiber photometry recordings, how to correct for calcium-independent artifacts, and important considerations for dual-color photometry imaging.

Recording the activity of a group of neurons in a freely-moving animal is a challenging undertaking. Moreover, as the brain is dissected into smaller and ...