The authors sincerely thank the Canadian Institutes of Health Research MOP 130258 and the Star Foundation for Children&#8217;s Health Research, along with the Molly Towell Foundation, for the provision of the research facility and financial support. The authors also sincerely thank Dr. Kinkead Richard for manuscript proofreading and advice.

This method required the use of animal subjects, allowed by Laval University Animal Ethics Committee (protocol # 2012-170).
1. Setup and Preparation

<ol>
	<li>Solutions
	<ol>
		<li>Prepare aCSF stock solutions according to the following recipes 7,17. Other recipes with concentration variations are available in the literature. Store stock solutions at 4 &#176;C for up to one month.
		<ol>
			<li>Salt solution: add 75.39 g of NaCl (129 mM final); 2.5 g of KCl (3.35 mM final); 0.81 g of NaH2PO4 (0.58 mM final); 2.33 g of MgCl2 (1.15 mM final); 1.85 g of CaCl2 (1.26 mM). Dissolve in 800 ml of distilled water then fill to 1 L with distillated water.</li>
			<li>Bicarbonate solution: add 17.65 g of NaHCO3 (21 mM final). Dissolve in 900 ml of distillated water then fill to 1 L with distilled water. Variations of the bicarbonate concentration will result in pH variations.</li>
			<li>Glucose solution: add 54.06 g of glucose (30 mM final). Dissolve in 400 ml of distillated water then fill to 500 ml with distilled water.</li>
		</ol>
		</li>
		<li>Prepare aCSF by diluting 100 ml of salt solution, 100 ml of bicarbonate solution, and 50 ml of glucose solution in 1 L of distilled water. Store at 4 &#176;C until use.</li>
		<li>Prepare a glass electrode by warming and stretching a glass tube until it breaks. Sand down the tip before use. The electrode can be reused many time as long as it stays clean.</li>
	</ol>
	</li>
	<li>Experimental Setup 
	Note: The set-up details are presented in Figure 2.
	<ol>
		<li>Turn on the amplifier, moving averager, data acquisition system, temperature controller and pump. Check the signal amplification (gain = 10,000), the filtration (low threshold, 10 Hz; high threshold, 5 kHz), and the sampling rate of the analog to digital conversion of the raw signal (2.5 kHz).</li>
		<li>Fill a bottle with aCSF and bubble it with carbogen (95% O2, 5% CO2). Induce a bubbled- and warmed-aCSF flow in the recording chamber (volume 5 ml) at 4 ml/min. Hold the temperature in the recording chamber at 26 (&#177; 1) &#176;C. pH should be 7.4 in these standard conditions.</li>
		<li>Keep 50 ml of RT and carbogen-bubbled aCSF in a 50 ml syringe near the dissection chamber.</li>
		<li>Turn on the computer and start the recording software.</li>
	</ol>
	</li>
</ol>
2. Dissection
<ol>
	<li>Weigh and visually determine the sex of the animal (males have a longer&#160;ano-genital distance, while females have a short&#160;ano-genital distance; males genitals are often dark while female genitals are pink).</li>
	<li>Anesthetize the newborn rodents by one of the following options: cryoanesthesia (completely immerse the animal in ice for 4 &#8211; 5 min 18), injection (equitensine at 4 ml/kg) 19 or inhalation of volatile anesthetics (isoflurane 20 or ether 21). Confirm an adequate plane of anesthesia by the absence of a paw withdrawal reflex.</li>
	<li>Place the animal on the bench, ventral face down. Section the rostral part of the head coronally with a scalpel at the bregma level (visible through the skin and skull). Perform this step immediately after the animal is anesthetized.</li>
	<li>Section the body coronally with a scalpel under the anterior members .</li>
	<li>Remove skin, muscles, fatty tissues and viscus with surgical and thin scissors and pliers. Since bones are soft at this age, be cautious to not damage the nervous system. Perform this step on the bench.</li>
	<li>Place the preparation in the dissection chamber. Use the aCSF stored in the syringe to oxygenate the preparation. From this point to the end of the recording, use a microscope.</li>
	<li>On the dorsal face of the preparation, cut the skull and vertebras from the rostral to the caudal part along the medium axis with surgical and thin scissors and pliers. Open the cut skull and vertebras in order to expose the nervous tissue. Use the aCSF strored in the syringe to oxygenate the preparation.</li>
	<li>With the pliers, remove the Arachnoid membrane, a thin tissue covering the surface of the nervous tissue. Retain the pia mater and blood vessels against the nervous tissue. Use the aCSF stored in the syringe to oxygenate the preparation.</li>
	<li>Place the preparation with the dorsal face down, and carefully bring down the brainstem and spinal cord by cutting the nerves and connective tissue with the scissors while holding the preparation in place. A dorsal approach is also possible. Keep the roots and nerves as long as possible. Remove the bones to isolate the brainstem and spinal cord. Use the aCSF stored in the syringe to oxygenate the preparation.</li>
	<li>Remove the cerebellum and remaining rostral structures by sectioning them with the scalpel. Use the aCSF stored in the needle to oxygenate the preparation.</li>
	<li>Optionally depending on experimental design, remove the pons with the scalpel by a coronal section anterior to the inferior cerebellar artery 22. Note that keeping the whole pons will slow down the rhythm in rats and fully inhibit it in mice. Preparations that include only the caudal part of the pons display a respiratory-like activity with a stable frequency at the C4 root.</li>
</ol>
3. Recording
<ol>
	<li>Place the preparation in the recording chamber, ventral face up. Fix the preparation with pins in the lowest part of the spinal cord and rostral-most part of the brainstem.</li>
	<li>Using a syringe linked to the electrode by its needle, induce a depression in the electrode (glass tip diameter: 150 - 225 &#181;m) by pulling out the syringe piston, in order to partially fill the electrode with aCSF.</li>
	<li>Using a micromanipulator, carefully place the electrode close to one of the fourth ventral roots. Other ventral roots could also present a respiratory-like activity (e.g., XII cranial nerve, C1 ventral root).</li>
	<li>Induce a depression in the electrode tank via the syringe by pulling out the piston in order to gently aspirate the nerve rootlet. Then carefully move the electrode to apply it against the spinal cord. 
	Note: Ideally the size of the rootlet matches the size of the electrode opening and thus creates a seal between the internal and external compartments of the electrode. Since differential amplifiers are commonly used for such recordings, any opening between the inside of the electrode and the recording chamber reduces the quality of the signal and makes it difficult to eliminate background noise.</li>
	<li>Start the recording.</li>
	<li>Record the rhythm produced by the preparation under normoxic conditions (i.e., aCSF bubbled with carbogen: 95% O2 and 5% CO2) for at least 20 min to determine the baseline parameters of the preparation.</li>
	<li>Switch the perfusion from carbogen-bubbled aCSF to stimulus aCSF (i.e., bubbled with 95% N2 and 5% CO2 for hypoxia stimulus) for 15 min. Alter exposure duration depending of the experimental protocol. Record the exposure duration on the recording and animal datasheet. Use stainless steel tubes between the aCSF bottle and the chamber whenever possible to avoid gas diffusion to the outside of the tube.</li>
	<li>Switch the perfusion back to standard carbogen-bubbled aCSF for at least 15 min for a recovery recording. Record this on the recording and animal datasheet.</li>
	<li>End the recording.</li>
</ol>
4. Statistical Analysis
<ol>
	<li>From the integrated signal, calculate the frequency as the number of bursts per minute (expressed in bursts/min), the amplitude as the difference between the baseline and the peak of the burst (expressed in mV), the burst duration as the duration from the beginning to the end of the burst (expressed in s), the burst area as the area under the curve of the burst on the integrated signal (expressed in mV&#183;s) (Figure 3).</li>
	<li>Calculate the frequency, amplitude, burst duration and burst area under normoxic (baseline), hypoxic (stimulus) and post-hypoxic (post-stimulus) recovery conditions as the average of the last 5 min of the recordings for each condition. Do not analyze the first portion of each condition because there is a delay between switching the perfusion and aCSF homogenization in the recording chamber.</li>
	<li>Express then stimulus (i.e., hypoxia) and post-stimulus (i.e., post-hypoxic) recovery values as a percentage of the baseline values for the corresponding recording. Express the results as means &#177; S.D.</li>
</ol>

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The authors have no competing financial interests or conflicts of interests to disclose.

Accurate quantification of respiratory activity can be challenging. Indeed, breathing is a function that can be both automatic and voluntary, and that is modulated according to the environment, the body&#8217;s needs, the emotional state and the behavior. The advantage of this technique is the isolation of the neural elements responsible for producing the respiratory command. Thus, electrophysiological recordings of brainstem-spinal cord preparations and plethysmography are complementary techniques to study the whole neu...

Breathing is a complex and vital activity controlled by the brain, allowing dioxygen (O2) uptake and carbon dioxide (CO2) elimination. The central respiratory drive is generated by a complex network located in the brainstem in both mammals 1, amphibians 2, reptiles 3, birds 4 and fishes 5. Even if the study of breathing can be processed in vivo, precise mechanistic investigations require direct access to the respiratory control network. To this end, Adrian and Buytendijk developed a reduced goldfish preparation, in which electrodes placed on the brainstem surface record the generated rhythm associated with gill ventilation 5. This approach was subsequently adapted by Suzue in 1984 6 for use in newborn rodents. The advent of this preparation has led to significant advances in respiratory neurobiology. Since it is relatively simple, the technique presented here is amenable to a broad range of basic investigations of rhythmic motor behaviors and their origins in newborn rodents.
The overall goal of this method is to record the neural correlate of inspiratory activity, a respiratory-like rhythm called fictive breathing, produced by the respiratory network. This method can be employed in a wide range of research objectives, targeting inspiratory responses to respiratory variations or pharmacology in both wild type 7 and transgenic 8 animals. Given that experiments are performed at a low temperature, without sensory afferents, and under conditions where the concentrations of glucose and O2 within the aCSF are high, questions have been raised regarding the physiological relevance of the signal recorded. While there are clear differences between in vivo and in vitro conditions (e.g., the frequency of inspiratory bursts) the fact remains that the presence of the core elements of the respiratory network 6 make it possible to study a robust rhythm associated to a vital homeostatic function 9,10.
The rationale behind the development and the use of this technique is to facilitate direct access to the brainstem elements of the respiratory network, which are hardly accessible in vivo, especially in newborns. The brainstem is placed under strictly controlled conditions: the recorded rhythm is not modulated by peripheral afferent inputs from the lungs or the carotid bodies, allowing the study to focus on the central respiratory drive itself 11. Thus, this access is utilized to apply stimuli and record the output signal. In contrast to plethysmography recordings, the respiratory rhythm is modulated by all of its components throughout the body (e.g., lung distension, peripheral chemosensors), making it difficult to apply precise stimuli.
In a newborn rat, the protocol consists of recording the fourth ventral root signal on an isolated brainstem and a truncated spinal cord, maintained in artificial cerebro-spinal fluid (aCSF). The rhythm generated by brainstem-spinal cord preparations is composed of individual slow bursts that are linked to the inspiratory signal 9. Isolated brainstem-spinal cord preparations are easily recordable in rats from post-natal day 0 to 4 (P0 - P4) 7. This approach is commonly used to evaluate the hypoxic response of the respiratory network, and also the response to hypercapnia, acidosis or drugs. An acute hypoxia protocol is presented here. This stimulation is obtained by withdrawal of O2 in the aCSF; this approach is commonly used to assess the tolerance and responsiveness to hypoxic insults. The protocol induces a rhythm depression from the first minute&#160;until the end of the hypoxia exposure (Figure 1)12. This depression is reversed during post-hypoxic recovery 12. Concerning the experimental design, it is important to notice that the pons, located at the rostral part of the brainstem, has an inhibitory action on the rhythm generator 8. Thus, preparations of full brainstem and rostral spinal cord display a lower rhythm. Inclusion of the pons in the isolated sample for the recording is determined according to the goal of the experiment 13; the study of pontine influence on the medulla oblongata network would require recordings with and without the pons to compare the results 14. Moreover, one of the advantages of this technique is the possibility of extending the rostral part of the preparation to include mesencephalic and/or diencephalic regions 15,16, making it possible to assess the effect of these regions on the ponto-medullary respiratory network.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Sylgard</td>
			<td>Sigma Aldrich</td>
			<td>761036-5EA</td>
			<td>Use under hood</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Bioshop</td>
			<td>SOD002</td>
			<td></td>
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			<td>KCl</td>
			<td>Bioshop</td>
			<td>POC888</td>
			<td></td>
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			<td>CaCl2</td>
			<td>Bioshop</td>
			<td>CCL444</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Bioshop</td>
			<td>MAG510</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Bioshop</td>
			<td>SOB999</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Bioshop</td>
			<td>SPM306</td>
			<td></td>
		</tr>
		<tr>
			<td>D-glucose</td>
			<td>Bioshop</td>
			<td>GLU501</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbogen</td>
			<td>Linde</td>
			<td>343-02-0006&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature Controller</td>
			<td>Warner Instruments, Hamden, CT, USA</td>
			<td>TC-324B</td>
			<td></td>
		</tr>
		<tr>
			<td>Suction electrode</td>
			<td>A-M Systems, Everett, WA, USA</td>
			<td>model 573000</td>
			<td></td>
		</tr>
		<tr>
			<td>Differential AC amplifier</td>
			<td>A-M Systems, Everett, WA, USA</td>
			<td>model 1700</td>
			<td></td>
		</tr>
		<tr>
			<td>Moving averager</td>
			<td>CWE, Ardmore, PA, USA</td>
			<td>model MA-821</td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquisition system</td>
			<td>Dataq Instruments, Akron, OH, USA</td>
			<td>model DI-720</td>
			<td></td>
		</tr>
		<tr>
			<td>LabChart software</td>
			<td>ADInstruments, Colorado Springs, CO, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Prism sofware</td>
			<td>Graphpad, La Jolla, CA, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection chamber</td>
			<td>Plastic box (e.g. petri box) will do</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Recording chamber</td>
			<td>Home made</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Base</td>
			<td>Kanetec, Bensenville, IL, USA</td>
			<td>MB</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>World Precision Instrument Inc, Sarasota, FL, USA</td>
			<td>KITE-R</td>
			<td></td>
		</tr>
		<tr>
			<td>Base</td>
			<td>Kanetec, Bensenville, IL, USA</td>
			<td>MB</td>
			<td></td>
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		<tr>
			<td>Peristaltic pump</td>
			<td>Gilson, Middleton, WI, USA</td>
			<td>MINIPULS 3</td>
			<td></td>
		</tr>
		<tr>
			<td>Faraday Cage</td>
			<td>Home made</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Computer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
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</table>

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.

As mentioned in the introduction, one of the most important advantages of this technique is the direct access to the brainstem to apply various stimuli. As an example, hypoxia was applied here. Figure 1.A.B. displays a full protocol recording, with both normoxic and hypoxic conditions. Figure 1.C.E. displays the rhythm recorded in normoxic conditions (i.e., aCSF bubbled with 95% O2 and 5% CO2 at 26 &#176;C). As previously demonstrated in th...

Watch this Scientific Journal Video about Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording at JoVE.com

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

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 ...

electrophysiology-on-isolated-brainstem-spinal-cord-preparations-from

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Neuroscience

8.5K vues.  Laval University. 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.

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

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

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 ...

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 ...

Neural Circuit Recording from an Intact Cockroach Nervous System

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological effects of insecticides. This article describes the scope of cockroach sensory modalities that can be used to assay how an insect nervous system responds to environmental perturbations. Emphasis here is on the escape behavior mediated by cerci to giant fiber transmission in Periplaneta americana. This in situ preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of neuronal activity. Peptides or other chemical reagents can then be applied directly to the nervous system in solution with the physiological saline. Insecticides could also be administered prior to dissection and the escape circuit can serve as a proxy for the excitable state of the central nervous system. In this context the assays described herein would also be useful to researchers interested in limb regeneration and the evolution of nervous system development for which P. americana is an established model organism.

This article describes the cockroach ventral nerve cord dissection and extracellular recordings from the cercal nerve and connectives. Evoked responses are generated by electrical stimulation of the cercal nerve or direct mechanical stimulation of the cerci.

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a signal driving the rhythmic contraction of inspiratory muscles. Rhythmic neural activity generated in the pBC and carried to other neuronal pools to drive the musculature of breathing may be studied using various approaches, including en bloc nerve recordings and transverse slice recordings. However, previously published methods have not extensively described the brainstem-spinal cord dissection process in a transparent and reproducible manner for future studies. Here, we present a comprehensive overview of a method used to reproducibly cut rhythmically-active brainstem slices containing the necessary and sufficient neuronal circuitry for generating and transmitting inspiratory drive. This work builds upon previous brainstem-spinal cord electrophysiology protocols to enhance the likelihood of reliably obtaining viable and rhythmically-active slices for recording neuronal output from the pBC, hypoglossal premotor neurons (XII pMN), and hypoglossal motor neurons (XII MN). The work presented expands upon previous published methods by providing detailed, step-by-step illustrations of the dissection, from whole rat pup, to in vitro slice containing the XII rootlets.

This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

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.

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 ...

Electromagnetic Controlled Closed-Head Model of Mild Traumatic Brain Injury in Mice

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and understanding the mechanisms of how a TBI alters brain function. The availability of multiple animal models of TBI is necessary to model the different aspects and severities of TBI seen in people. This manuscript describes the use of a midline closed head injury (CHI) to develop a mouse model of mild TBI. The model is considered mild because it does not produce structural brain lesions based on neuroimaging or gross neuronal loss. However, a single impact creates enough pathology that cognitive impairment is measurable at least 1 month after injury. A step-by-step protocol to induce a CHI in mice using a stereotaxically guided electromagnetic impactor is defined in the paper. The benefits of the mild midline CHI model include the reproducibility of the injury-induced changes with low mortality. The model has been temporally characterized up to 1 year after the injury for neuroimaging, neurochemical, neuropathological, and behavioral changes. The model is complementary to open skull models of controlled cortical impact using the same impactor device. Thus, labs can model both mild diffuse TBI and focal moderate-to-severe TBI with the same impactor.

The protocol describes mild traumatic brain injury in a mouse model. In particular, a step-by-step protocol to induce a mild midline closed head injury and the characterization of the animal model is fully explained.

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and ...