microelectrode array-based assessment of neuronal networks in mouse spinal cord slices

Microelectrode Array-Based Assessment of Neuronal Networks in Mouse Spinal Cord Slices

To begin recording dorsal horn activity, transfer the slice from the incubator to the MEA well using a large-tip tip Pasteur pipette filled with artificial CSF, and add additional artificial CSF.
Use a fine, short-hair paintbrush to position the slice over the 60-electrode recording array. Then, place a weighted net 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.

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All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1.&#160;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 incubation chamber, where slices are stored until recording commences and during experiments as both perfusate and diluent for drugs. See Table 1 for the detailed composition.</li>
	</ol>
	</li>
	<li>Prepare aCSF containing (in mM) 118 NaCl, 25 NaHCO3, 10 glucose, 2.5 KCl, 1 NaH2PO4, 1 MgCl2, and 2.5 CaCl2 by adding the required quantities of the above, excluding CaCl2, to 2 L of distilled water.</li>
	<li>Bubble the above solution with carbogen (95% O2, 5% CO2) for 5 min and add CaCl2. 
	NOTE: This step prevents CaCl2 precipitation, i.e., the solution should not turn cloudy. For drug application during experiments, dilute the drug stock solutions in aCSF to desired final concentrations.</li>
</ol>
2. Sucrose-substituted artificial cerebrospinal fluid
NOTE: Sucrose-substituted aCSF is used during dissection and spinal cord slicing. As indicated by the name, sucrose is substituted for NaCl to reduce neuronal excitation during these procedures while maintaining osmolarity. See Table 1 for the detailed composition.
<ol>
	<li>Prepare sucrose-substituted aCSF containing (in mM) 250 sucrose, 25 NaHCO3, 10 glucose, 2.5 KCl, 1 NaH2PO4, 1 MgCl2, and 2.5 CaCl2 by adding the required quantities of all of the above, excluding CaCl2, to 300 mL of distilled water.</li>
	<li>Bubble the solution with carbogen for 5 min and then add CaCl2.</li>
	<li>Store the solution in a -80 &#176;C freezer for approximately 40 min or until the solution forms a slurry. Avoid freezing solid and use while in slurry consistency.</li>
</ol>
3. Microelectrode array preparation
NOTE: The contact surface of the MEA requires a pretreatment to make it hydrophilic.
<ol>
	<li>Before the experiment, fill the MEA well with either fetal bovine serum (FBS) or horse serum (HS) for 30 min.</li>
	<li>Remove the FBS or HS and thoroughly rinse MEA with approximately five washes of distilled water until the distilled water is no longer foamy. Fill the well with aCSF, ready for use.</li>
</ol>
4. Acute spinal cord slice preparation
<ol>
	<li>Deeply anesthetize the mouse with 100 mg/kg ketamine (i.p.) and then decapitate it using large surgical scissors.</li>
	<li>Remove the skin over the abdominal region by making a small cut in the skin at the level of the hips. Pull the skin on either side of the cut rostrally until all the skin is removed, i.e., from the top of the rib cage to the top of the pelvis (both ventrally and dorsally).</li>
	<li>Place the body on ice and use a ventral approach to expose the vertebral column by removing all the viscera and cutting through the ribs lateral to the sternum.</li>
	<li>Remove the ventral rib cage, both scapulae (cut off at approximately T2), and the lower limbs and pelvis (cut off at approximately the top of the sacrum).</li>
	<li>Transfer the vertebral column and rib preparation to a dissecting bath containing ice-cold sucrose aCSF. Pin all four corners of the preparation (ventral surface upwards) by placing pins through the lower back muscles and the attached upper ribs.</li>
	<li>Remove all muscle and connective tissue overlying the ventral surface of the vertebrae with rongeurs and identify the vertebral region over the lumbosacral enlargement, which lies approximately beneath the T12 to L2 vertebral bodies.</li>
	<li>Remove a vertebral body that is caudal to the lumbosacral enlargement region to provide access to the spinal cord as it sits in the vertebral canal.</li>
	<li>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.</li>
	<li>Once the vertebral bodies are removed to reveal the lumbosacral enlargement, carefully clear the remaining roots that anchor the spinal cord with spring scissors until the cord floats free.</li>
	<li>Isolate the spinal cord with rostral and caudal cuts well above and below the lumbosacral enlargement, allowing the target region of the cord to &#39;float free.&#39; 
	NOTE: The preferred slice orientation will determine how the cord is subsequently mounted for sectioning (Figure 1).</li>
	<li>For transverse slices, lift the lumbosacral segment by an attached root and place it on a pre-cut polystyrene (Styrofoam) block (1 cm x 1 cm x 1 cm) with a shallow channel cut in the center. Use cyanoacrylate adhesive (see the Table of Materials) to attach the block and cord to the sectioning platform and place it in the cutting bath containing ice-cold sucrose aCSF (slurry). 
	NOTE: The shallow channel helps secure and orient the spinal cord, with the dorsal side exposed and the thoracic end of the cord at the bottom of the block.</li>
	<li>For sagittal slices, lay a thin line of cyanoacrylate adhesive on the sectioning platform, lift the lumbosacral enlargement by an attached root, and place the cord along the line of glue, ensuring one lateral surface is in the adhesive and the other faces upwards. Place it in the cutting bath containing ice-cold sucrose aCSF (slurry).</li>
	<li>For horizontal slices, put a thin line of cyanoacrylate adhesive on the sectioning platform. Lift the lumbosacral enlargement by an attached root, and place the lumbosacral enlargement along the line of adhesive, ensuring the ventral surface is in the adhesive and the dorsal surface faces upwards. Use attached roots to position the cord. Place it in the cutting bath containing ice-cold sucrose aCSF (slurry).</li>
</ol>
5. Microelectrode array recordings
NOTE: The following steps detail how to use record data from MEA-based experiments on spinal cord slices. Several MEA designs can be used depending on the experiment. Design details for MEAs used in these experiments are shown in Table 2 and Figure 2. Detailed design information has been published by Egert et al. and Thiebaud et al. for planar and 3-dimensional (3D) MEAs, respectively. Both MEA types are composed of 60 titanium nitride electrodes, with a silicon nitride insulating layer and titanium nitride tracks and contact pads.
<ol>
	<li>Experimental setup
	<ol>
		<li>Turn on the computer and interface board, and start the recording software.</li>
		<li>Load the pre-assembled recording template (Figure 3A). Name the files for the day in the recorder tab.</li>
		<li>Continuously bubble aCSF with carbogen (5% CO2, 95% O2) for the duration of the experiment.</li>
		<li>Turn the perfusion system on, which is controlled by a peristaltic pump. Place the inlet line into aCSF and the inlet end in a waste beaker. Prime the perfusion lines with aCSF.</li>
		<li>Prepare 4-AP and any other drug solutions by diluting stocks in 50 mL of aCSF to the required final concentration (e.g., 200 &#181;M for 4-AP).</li>
	</ol>
	</li>
	<li>4-aminopyridine (4-AP) activity
	<ol>
		<li>Following incubation, transfer a single slice from the incubator using a large-tip Pasteur pipette filled with aCSF.</li>
		<li>Place the slice in the MEA well and add additional aCSF.</li>
		<li>Position the slice over the 60-electrode recording array using a fine short hair paintbrush. Avoid contacting the electrodes with the paintbrush or dragging the tissue across the electrodes, especially if using 3D arrays. 
		NOTE: Depending on the MEA layout, this can be done with or without the assistance of a microscope for accurate positioning.</li>
		<li>After positioning the slice, place a weighted net over the tissue to hold it in place and promote good contact with MEA electrodes. 
		NOTE: The slice may need repositioning following net placement.</li>
		<li>Place the MEA in the recording headstage (Figure 2A, B).</li>
		<li>Check the position of the tissue over the electrodes using an inverted microscope (2x magnification) to confirm that as many electrodes as possible are under the superficial dorsal horn (SDH). Ensure that at least 2-6 electrodes do not contact the slice as these electrodes are important for subtracting noise and recording artefacts during analysis (Figure 2E).</li>
		<li>Turn on the camera, connect it to the device, and take a reference image of the slice relative to the MEA for use during analysis.</li>
		<li>Press Start DAQ (data acquisition) in the recording software and confirm that all electrodes are receiving a clear signal. 
		NOTE: If the signal is noisy, unclip the headstage, and clean both the MEA contact pads and gold spring contacts with 70% ethanol (use a laboratory wipe to ensure that the pads and contacts are dry after cleaning). If the signal is still noisy, turn off the malfunctioning electrodes in the recording software or note down for exclusion later during analysis.</li>
		<li>Attach the perfusion inlet and outlet lines to the MEA-well (previously filled with aCSF) and turn the perfusion system on. Check the flow rate, ideally 4-6 bath volumes per minute, and ensure that the outflow is sufficient to prevent overflow of the superfusate.</li>
		<li>Allow the tissue to equilibrate for 5 min and then record 5 min of raw, unfiltered baseline data.</li>
		<li>Move the perfusion inlet line from aCSF to a 4-AP solution and wait for 12 min for the 4-AP-induced rhythmic activity to reach steady state (2 min for drugs to reach the bath and 10 min for the activity to peak and then plateau).</li>
		<li>Record 5 min of 4-AP-induced activity. Be prepared for subsequent recordings to test the drugs or to check the stability of 4-AP.</li>
	</ol>
	</li>
</ol>
Table 1: Artificial Cerebrospinal Fluid compositions. 
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemical</td>
			<td>aCSF (mM)</td>
			<td>aCSF (g/100 mL)</td>
			<td>Sucrose-substituted aCSF (mM)</td>
			<td>Sucrose-substituted aCSF (g/100 mL)</td>
			<td>High-potassium aCSF (mM)</td>
			<td>High-potassium aCSF (g/100 mL)</td>
		</tr>
		<tr>
			<td>Sodium chloride (NaCl)</td>
			<td>118</td>
			<td>0.690</td>
			<td>-</td>
			<td>-</td>
			<td>118</td>
			<td>0.690</td>
		</tr>
		<tr>
			<td>Sodium hydrogen carbonate (NaHCO3)</td>
			<td>25</td>
			<td>0.210</td>
			<td>25</td>
			<td>0.210</td>
			<td>25</td>
			<td>0.210</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>10</td>
			<td>0.180</td>
			<td>10</td>
			<td>0.180</td>
			<td>10</td>
			<td>0.180</td>
		</tr>
		<tr>
			<td>Potasium chloride (KCl)</td>
			<td>2.5</td>
			<td>0.019</td>
			<td>2.5</td>
			<td>0.019</td>
			<td>4.5</td>
			<td>0.034</td>
		</tr>
		<tr>
			<td>Sodium dihydrogen phosphate (NaH2PO4)</td>
			<td>1</td>
			<td>0.012</td>
			<td>1</td>
			<td>0.012</td>
			<td>1</td>
			<td>0.012</td>
		</tr>
		<tr>
			<td>Magnesium cloride (MgCl2)</td>
			<td>1</td>
			<td>0.01</td>
			<td>1</td>
			<td>0.01</td>
			<td>1</td>
			<td>0.01</td>
		</tr>
		<tr>
			<td>Calcium chloride (CaCl2)</td>
			<td>2.5</td>
			<td>0.028</td>
			<td>2.5</td>
			<td>0.028</td>
			<td>2.5</td>
			<td>0.028</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>-</td>
			<td>-</td>
			<td>250</td>
			<td>8.558</td>
			<td>-</td>
			<td>-</td>
		</tr>
	</tbody>
</table>
Table 2: Microelectrode array layouts.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="5">Microelectrode Array Layouts</td>
		</tr>
		<tr>
			<td>Microelectrode Array Model</td>
			<td>60MEA 200/30iR-Ti</td>
			<td>60-3DMEA 100/12/40iR-Ti</td>
			<td>60-3DMEA 200/12/50iR-Ti</td>
			<td>60MEA 500/30iR-Ti</td>
		</tr>
		<tr>
			<td>Planar or 3-Dimensional (3D)</td>
			<td>Planar</td>
			<td>3D</td>
			<td>3D</td>
			<td>Planar</td>
		</tr>
		<tr>
			<td>Electrode Grid</td>
			<td>8 x 8</td>
			<td>8 x 8</td>
			<td>8 x 8</td>
			<td>6 x 10</td>
		</tr>
		<tr>
			<td>Electrode Spacing</td>
			<td>200 &#181;m</td>
			<td>100 &#181;m</td>
			<td>200 &#181;m</td>
			<td>500 &#181;m</td>
		</tr>
		<tr>
			<td>Electrode Diameter</td>
			<td>30 &#181;m</td>
			<td>12 &#181;m</td>
			<td>12 &#181;m</td>
			<td>30 &#181;m</td>
		</tr>
		<tr>
			<td>Electrode Height (3D)</td>
			<td>N/A</td>
			<td>40 &#181;m</td>
			<td>50 &#181;m</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Experiments</td>
			<td>Transverse slice</td>
			<td>Transverse slice</td>
			<td>Sagittal + Horizontal</td>
			<td>Sagittal + Horizontal</td>
		</tr>
	</tbody>
</table>

<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&#160;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>

Source: Iredale, J. A., et al. <a href="https://app.jove.com/v/62920">Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays.</a> J. Vis. Exp. (2022).
This video demonstrates a microelectrode array-based assay for studying neuronal network activity in mouse spinal cord sections. First, the electrophysiological activity from the superficial dorsal horn (SDH) of the slice is recorded. Then, a potassium channel inhibitor is introduced to prolong depolarization, resulting in synchronous rhythmic activity across the neuronal network.

<img alt="Figure 1" src="/files/ftp_upload/22746/22746_Figure1.jpg" /> 
Figure 1: Spinal cord slice orientations, mounting and cutting methods. (A) Transverse slices require a Styrofoam cutting block with a supporting groove cut into it. The spinal cord is rested against the block in the support groove, the dorsal side of the cord facing away from the block. The block and cord are glued onto a cutting stage with cyanoacrylate adhesive. (B</st...

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1.&#160;In vitro electrophysiology

	...

Take a microelectrode array, or MEA, well, containing aCSF. Position a mouse spinal cord slice in the well and secure it with a weighted net.
Place the MEA well in a recording system. Under a microscope, ensure maximal contact between the MEA electrodes and the superficial dorsal horn or SDH, a region rich in interneurons.
Maintain a continuous aCSF flow to equilibrate the tissue.
Neurons communicate through action potentials. Sodium ion influx depolarizes the membrane, followed by potassium ion outflow that causes repolarization. The membrane briefly hyperpolarizes, preventing a new action potential until the resting potential is restored.
Record spontaneous neuronal activity from the MEA electrodes. Signals co-occurring across multiple electrodes indicate a network of synaptically linked neurons.
Introduce a potassium channel inhibitor to prolong depolarization that leads to increased action potential frequency and synchronous rhythmic activity across the network.
More electrodes showing coincident signals indicate the inhibitor-mediated synchronous activity.

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视频: 基于微电极阵列的小鼠脊髓切片神经元网络评估 - 实验

Simultaneous Recordings of Cortical Local Field Potentials and Electrocorticograms in Response to Nociceptive Laser Stimuli from Freely Moving Rats

Electrocortical responses, elicited by laser heat pulses that selectively activate nociceptive free nerve endings, are widely used in many animal and human studies to investigate the cortical processing of nociceptive information. These laser-evoked brain potentials (LEPs) consist of several transient responses that are time-locked to the onset of laser stimuli. However, the functional properties of the LEP responses are still largely unknown, due to the lack of a sampling technique that can simultaneously record neural activities at the surface of the cortex (i.e., electrocorticogram [ECoG] and scalp electroencephalogram [scalp EEG]) and inside the brain (i.e., local field potential [LFP]). To address this issue, we present here an animal protocol using freely moving rats. This protocol is composed of three main procedures: (1) animal preparation and surgical procedures, (2) a simultaneous recording of ECoG and LFP in response to nociceptive laser stimuli, and (3) data analysis and feature extraction. Specifically, with the help of a 3D-printed protective shell, both ECoG and LFP electrodes implanted on the rat&#39;s skull were securely held together. During data collection, laser pulses were delivered on the rat&#39;s forepaws through gaps in the bottom of the chamber when the animal was in spontaneous stillness. Ongoing white noise was played to avoid the activation of the auditory system by the laser-generated ultrasounds. As a consequence, only nociceptive responses were selectively recorded. Using the standard analytical procedures (e.g., band-pass filtering, epoch extraction, and baseline correction) to extract stimulus-related brain responses, we obtained results showing that LEPs with a high signal-to-noise ratio were simultaneously recorded from ECoG and LFP electrodes. This methodology makes the simultaneous recording of ECoG and LFP activities possible, which provides a bridge of electrocortical signals at the mesoscopic and macroscopic levels, thereby facilitating the investigation of nociceptive information processing in the brain.

We developed a technique that simultaneously records both electrocorticography and local field potentials in response to nociceptive laser stimuli from freely moving rats. This technique helps establish a direct relationship of electrocortical signals at the mesoscopic and macroscopic levels, which facilitates the investigation of nociceptive information processing in the brain.

在自由运动大鼠的反应中, 皮质局部电位和电皮质图谱的同时记录

Electrocortical responses, elicited by laser heat pulses that selectively activate nociceptive free nerve endings, are widely used in many animal and ...

科研

JoVE Journal

行为学

Objective Nociceptive Assessment in Ventilated ICU Patients: A Feasibility Study Using Pupillometry and the Nociceptive Flexion Reflex

The concept of objective nociceptive assessment and optimal pain management have gained increasing attention. Despite the known negative short- and long-term consequences of unresolved pain or excessive analgosedation, adequate nociceptive monitoring remains challenging in non-communicative, critically ill adults. In the intensive care unit (ICU), routine nociceptive evaluation is carried out by the attending nurse using the Behavior Pain Scale (BPS) in mechanically ventilated patients. This assessment is limited by medication use (e.g., neuromuscular blocking agents) and the inherent subjective character of nociceptive evaluation by third parties.
Here, we describe the use of two nociceptive reflex testing devices as tools for objective pain evaluation: the pupillary dilation reflex (PDR) and nociception flexion reflex (NFR). These measurement tools are non-invasive and well tolerated, providing clinicians and researchers with objective information regarding two different nociceptive processing pathways: (1)&#160;the pain-related autonomic reactivity&#160;and (2) the ascending component of the somatosensory system. The use of PDR and NFR measurements are currently limited to specialized pain clinics and research institutions because of impressions that these are technically demanding or time-consuming procedures, or even because of a lack of knowledge regarding their existence.
By focusing on the two abovementioned nociceptive reflex assessments, this study evaluated their feasibility as a physiological pain measurement method in daily practice. Pursuing novel technologies for evaluating the analgesia level in unconscious patients may further improve individual pharmacological treatment and patient related outcome measures. Therefore, future research must include large well-designed clinical trials in a real-life environment.

Pain assessment in anesthetized patients who cannot communicate with the outside world in any way remains challenging despite the development of innovative objective pain evaluation tools. In this project, the pupillary dilation reflex and the nociception flexion reflex are assessed in critically ill, mechanically ventilated adult patients.

应用 Pupillometry 和伤害性屈曲反射的可行性研究--通气 ICU 患者的客观伤害评估

The concept of objective nociceptive assessment and optimal pain management have gained increasing attention. Despite the known negative short- and ...

医学

Whole-Brain 3D Activation and Functional Connectivity Mapping in Mice using Transcranial Functional Ultrasound Imaging

Functional ultrasound (fUS) imaging is a novel brain imaging modality that relies on the high-sensitivity measure of the cerebral blood volume achieved by ultrafast doppler angiography. As brain perfusion is strongly linked to local neuronal activity, this technique allows the whole-brain 3D mapping of task-induced regional activation as well as resting-state functional connectivity, non-invasively, with unmatched spatio-temporal resolution and operational simplicity. In comparison with fMRI (functional magnetic resonance imaging), a main advantage of fUS imaging consists in enabling a complete compatibility with awake and behaving animal experiments. Moreover, fMRI brain mapping in mice, the most used preclinical model in Neuroscience, remains technically challenging due to the small size of the brain and the difficulty to maintain stable physiological conditions. Here we present a simple, reliable and robust protocol for whole-brain fUS imaging in anesthetized and awake mice using an off-the-shelf commercial fUS system with a motorized linear transducer, yielding significant cortical activation following sensory stimulation as well as reproducible 3D functional connectivity pattern for network identification.

This protocol describes the quantification of volumetric cerebral hemodynamic variations in the mouse brain using functional ultrasound (fUS). Procedures for 3D functional activation map following sensory stimulation as well as resting-state functional connectivity are provided as illustrative examples, in anesthetized and awake mice.

使用经颅功能超声成像的小鼠全脑3D激活和功能连接映射

Functional ultrasound (fUS) imaging is a novel brain imaging modality that relies on the high-sensitivity measure of the cerebral blood volume achieved by ...

Microelectrode Array-Based Assessment of Neuronal Networks in Mouse Spinal Cord Slices

成績單

Microelectrode Array-Based Assessment of Neuronal Networks in Mouse Spinal Cord Slices