Name: simultaneous pre- and post-synaptic electrophysiological recording from xenopus nerve-muscle co-cultures
Uploaded: 2013-03-11T12:00:00
Duration: 8 min 13 s
Description: Watch this Scientific Journal Video about Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures at JoVE.com

Funded by the NSF (0854551).

1. Pre-experimental Preparation
<ol>
	<li>Prepare the following solutions (see Table 1 for compositions): (a) 1 liter NFR (Normal Frog Ringer), (b) 100 ml 10% saline, (c) 100 ml CMF (Ca2+ /Mg2+-free solution), (d) 1 ml ITS (Insulin-Transferrin-Selenium, Sigma I1884), (e) 100 ml L-15 culture medium, (f) 10 ml HCG (human chorionic gonadotropin Sigma CG-10), (g) 100 ml K+-internal solution, (h) 100 ml K+-internal solution for Amphotericin B. The osmotic strength of solution 1 should be checked to ensure that it is approximately 260 mOsM using a vapor pressure osmometer (e.g. Wescor model #5100C).</li>
	<li>Filter sterilize solutions (b) and (c). Solutions (a) (b) and (c) can be stored up to three months at 4 &deg;C. Prepare solution (d) by adding 1 ml deionized H2O to the vial containing the lyophilized powder via a 25 gauge syringe needle and syringe filter. This final solution can be stored at 4 &deg;C for 30 days.</li>
	<li>Prepare solution (e) in a laminar flow hood by combining all components in a beaker or flask. Filter sterilize the final solution and transfer 10 ml aliquots into sterile centrifuge tubes. Store at -20 &deg;C.</li>
	<li>Prepare the HCG (solution (f)) by adding 10 ml deionized H20 to the vial containing the lyophilized powder via a 25 gauge syringe needle and syringe filter. This final solution can be stored at 4 &deg;C for 30 days.</li>
	<li>Fabricate at least two microdissection tools by gluing a Minutien pin (26002-10, Fine Science Tools) to the end of a glass Pasteur pipette with cyanoacrylate glue. Allow the sharp end of the pin to extend past the end of the pipette approximately 0.5 cm.</li>
	<li>Two days before preparing cultures, induce breeding with the following procedure. Identify a breeding-ready pair of Xenopus by observing a prominent, reddish cloaca on the female and dark pigmentation on the planar surface of the front paws of the male. One at a time, net each frog and hold it ventral side down on a sink with the net so that it cannot escape. Inject 1 ml of solution (f) through the net and subcutaneously into one of the dorsal lymph sacs.</li>
	<li>Place the breeding pair together in a covered ten gallon tank of water. To ensure that the animals will not trample the newly laid and fertilized eggs, install a screened floor with a mesh size of approximately &frac12;" fitted approximately 1-2 inches above the bottom of the tank (Figure 1A).</li>
	<li>Leave frogs undisturbed for 12-48 hr until the animals are in amplexus and fertilized eggs are observed on the floor below the screen (Figure 1B).</li>
	<li>Remove the animals from the tank, but leave the eggs undisturbed for at least 24 hr longer. This pair of frogs can be re-bred after six weeks.</li>
	<li>Loosen the embryos from the bottom of the tank and transfer them to four or five 60x15 mm culture dishes containing 10% saline (solution b). Sort the embryos by stage according to the scheme of Niewkoop and Faber (ref. 20). Useful embryos will be those at stage 22-24. Figure 2A shows an embryo at approximately stage 22 while Figure 2B shows an embryo that is too far along in development to be useful (approximately stage 28). It is important to choose embryos that are healthy: those that are smooth in appearance with light brown and white mottling are ideal. Embryos that have large black or white patches are generally unhealthy and unusable.</li>
</ol>
2. Microdissection of Xenopus Embryos
<ol>
	<li>Inside a laminar flow hood, label and fill approximately halfway three 60x15 mm sterile culture dishes with 10 % saline; and one with CMF.</li>
	<li>Using a sterile, glass Pasteur pipette transfer five to ten stage 22-24 embryos into one of the dishes containing 10% saline. With the aid of a stereo zoom dissecting microscope inside the hood (0.6-5x with 10 x eyepieces), remove the jelly coat and vitelline membrane from each embryo using two pairs of sterile #5 forceps (11251-30, Fine Science Tools). (See Figures 3A and 3B.)</li>
	<li>Wash the bare embryos by passing them, one at a time, through the remaining two dishes of 10% saline and finally into the dish containing CMF. Use a new, sterile pipette for each transfer and minimize the volume of solution transferred from dish to dish.</li>
	<li>In turn, hold each embryo gently but firmly with a pair of forceps and, using the microdissection tool fashioned in step 1.5, remove the neural tube and associated myotomes which are located at the most dorsal aspect of the animal. Do this by making three slices, one at either end of the location of the neural tube and a third just ventral to it (Figure 4A). Move each dissected neural tube/myotome to a clean part of the dish away from the yolk granules and other debris (Figure 4B). The dorsal-ventral axis, and the locations of the neural tube and myotomes are indicated in Figure 4.</li>
	<li>After about fifteen minutes in this solution (CMF) use forceps to lift the pigmented skin free of the dissected tissue and discard. After an additional 30-60 min, the cells will form a "sand pile" as they become dissociated from one another (Figure 4C).</li>
</ol>
3. Preparation of Nerve-muscle Co-cultures
<ol>
	<li>Thaw a 10 ml tube of culture medium and aseptically add 70 &mu;l ITS and 35 ng/ml BDNF (Sigma B3795).</li>
	<li>Label and fill sterile 35 mm culture dishes (FD35-100 World Precision Instruments) approximately halfway with L-15 culture medium (solution (e)), one for each embryo dissected.</li>
	<li>Fabricate a plating pipette by grasping each end of a glass Pasteur pipette while holding the tapered portion over a flame. Pull the ends apart to about 10 cm and then break off the end to yield a tip of approximately 0.2 mm.</li>
	<li>Use the plating pipette to suck up the "sand pile" from one embryo minimizing the amount of solution drawn. Expel the cells onto the bottom of a culture dish. Plate the cells into several lines. Observe plated undifferentiated cells (Figure 5A and 5B).</li>
	<li>Leave the dishes of plated cells undisturbed for at least fifteen minutes to allow the cells time to attach to the dish.</li>
</ol>
4. Patch Clamping Nerve-muscle Synapses
<ol>
	<li>After 12-24 hr in culture, plated cells take on distinct morphological characteristics: muscle cells become spindle-shaped and spinal neurons extend long processes (Figure 6). Functional synaptic contact between neurite varicosities and muscle cells can be confirmed with simultaneous paired electrophysiological recordings. "M", "S" and "V" refer respectively to the muscle cell, neuronal soma and presynaptic varicosity.</li>
	<li>Prepare two patch electrodes for recording: one for the presynaptic varicosity and one for the muscle cell.</li>
	<li>For the presynaptic electrode, prepare an amphotericin-containing solution as follows: Add 100 &mu;l DMSO to a 1.5 ml microcentrifuge tube containing 5 mg amphotericin B (Sigma A4888). Vortex this solution for 10 sec. Next, add 10 &mu;l of this solution to a second microcentrifuge tube that contains 625 &mu;l K+-internal solution for Amphotericin B (solution (h)). Vortex as above.</li>
	<li>Fill the pre-synaptic electrode with the amphotericin-containing K+-internal solution prepared in step 4.3 and the post-synaptic electrode with K+-internal solution (solution (g)).</li>
	<li>Replace media in the culture dish with NFR and relocate it to an inverted microscope fitted with phase contrast optics.</li>
	<li>Position both electrodes just above their respective targets. Obtain the whole-cell configuration with the muscle cell electrode before the perforated patch configuration with the varicosity electrode.</li>
	<li>To confirm functionality of the synapse, depolarize the varicosity with a patch clamp amplifier (e.g. Axopatch 200B, Molecular Devices) under software control (e.g. by pCLAMP 9, Molecular Devices) and observe the simultaneous presynaptic and postsynaptic currents. Figure 7 shows the currents seen in response to depolarizing steps of voltage given to the presynaptic varicosity in 10 mV increments from -30 mV to +40 mV. The inward currents seen in the presynaptic cell are carried by Na+ and Ca2+ and the outward currents by K+. Currents recorded in the postsynaptic cell are responses to neurotransmitter released from the varicosity and are carried mostly by Na+ through nicotinic acetylcholine receptor channels.</li>
</ol>

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No conflicts of interest declared.

Key steps in the successful co-culturing of motoneurons and muscle cells are the use of appropriately staged embryos produced from the induced breeding of Xenopus frogs, and the careful aseptic dissection of the undifferentiated spinal neurons and myoblasts. Fertilized eggs should be left undisturbed until they reach approximately stage 10 or so as moving them sooner often halts their development. Healthy embryos are identified by a smooth appearance and a brown and white mottled coloring. Stage 22-24 embryos are the most useful because this is the point during development just after the neural tube closes and before the myocytes have differentiated significantly. In addition, cells from older embryos fail to dissociate well in CMF. Care should be taken when removing the vitelline membrane as it adheres strongly to the embryo. The membrane should be torn apart using sharp forceps so that the embryo emerges intact. Another important precaution is to carefully place the cells onto the bottom of the culture dish (rather than letting them settle there). This method is preferred as this increases the likelihood that the cells will adhere to the culture dish.
Functional neurotransmission between nerve and muscle can be determined with just postsynaptic recording and often can be ascertained by the observation of spontaneous muscle contraction after innervation. Un-innervated muscle cells do not twitch in culture. Postsynaptic recording alone is useful for recording miniature endplate currents or potentials but measurements of evoked release requires presynaptic stimulation, and correlation of pre- and postsynaptic currents requires double patch-clamp.
Besides the paired-patch recording method described here, this preparation offers the opportunity to introduce a third pipette at the neuronal soma5 This allows the generation of an action potential that can propagate to the presynaptic varicosity and lead to the release of neurotransmitter. In addition, putative agents that are thought to mediate or modulate synaptic transmission can be introduced to either side of the synapse: via the muscle cell pipette or through diffusion from a third pipette placed at the soma.

<table border="1">
 <tbody>
 <tr>
 <td>Equipment/Supplies</td>
 <td>Vendor</td>
 <td>Catalogue/Model Number</td>
 <td></td>
 </tr>
 <tr>
 <td>Insulin-Transferrin-Selenium</td>
 <td>Sigma</td>
 <td>I1884</td>
 <td></td>
 </tr>
 <tr>
 <td>Human Chorionic Gonadotropin </td>
 <td>Sigma</td>
 <td>CG-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Vapor Pressure Osmometer</td>
 <td>Wescor</td>
 <td>5100C</td>
 <td></td>
 </tr>
 <tr>
 <td>Minutien Pins</td>
 <td>Fine Science Tools</td>
 <td>26002-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Amphotericin B</td>
 <td>Sigma</td>
 <td>A4888</td>
 <td></td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>Fine Science Tools</td>
 <td>11251-30</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>
 <table border="1">
 <tbody>
 <tr>
 <td>Solution Name</td>
 <td>Recipe</td>
 </tr>
 <tr>
 <td>(a) NFR (Normal Frog Ringer (in mM)</td>
 <td>116 NaCl, 2 KCl, 1.8 CaCl2, 5 Na+-HEPES, pH 7.35.</td>
 </tr>
 <tr>
 <td>(b) 10% saline</td>
 <td>10 ml NFR+90 ml deionized H2O</td>
 </tr>
 <tr>
 <td>(c) CMF (Ca2+ /Mg2+-free solution)</td>
 <td>125 NaCl, 2 KCl, 1.2 EDTA, 5 Na+-HEPES, pH 7.35.</td>
 </tr>
 <tr>
 <td>(e) L-15 Culture Media</td>
 <td>50 ml L-15 media (Sigma L1518) + 50 ml NFR.</td>
 </tr>
 <tr>
 <td>(g) K+-internal solution (in mM)</td>
 <td>116 KCl, 1 NaCl, 1 MgCl2, 10 EGTA, 5 HEPES, pH 7.3</td>
 </tr>
 <tr>
 <td>(h) K+-internal solution for Amphotericin B (in mM) </td>
 <td>52 K2SO4, 38 KCl, 1 EGTA, 5 HEPES, pH 7.3</td>
 </tr>
 </tbody>
</table>
 </td>
 </tr>
 </tbody>
</table>
Table 1.

simultaneous pre- and post-synaptic electrophysiological recording from xenopus nerve-muscle co-cultures

Much information about the coupling of presynaptic ionic currents with the release of neurotransmitter has been obtained from invertebrate preparations, most notably the squid giant synapse1. However, except for the preparation described here, few vertebrate preparations exist in which it is possible to make simultaneous measurements of neurotransmitter release and presynaptic ionic currents. Embryonic Xenopus motoneurons and muscle cells can be grown together in simple culture medium at room temperature; they will form functional synapses within twelve to twenty-four hours, and can be used to study nerve and muscle cell development and synaptic interactions for several days (until overgrowth occurs). Some advantages of these co-cultures over other vertebrate preparations include the simplicity of preparation, the ability to maintain the cultures and work at room temperature, and the ready accessibility of the synapses formed2-4. The preparation has been used widely to study the biophysical properties of presynaptic ion channels and the regulation of transmitter release5-8. In addition, the preparation has lent itself to other uses including the study of neurite outgrowth and synaptogenesis9-12, molecular mechanisms of neurotransmitter release13-15, the role of diffusible messengers in neuromodulation16,17, and in vitro synaptic plasticity18-19.

Figure 4B shows the dorsal view of an isolated spinal cord/myotome immediately after its removal from a stage 22 Xenopus embryo. Figure 4C shows that, after skin removal and incubation in Ca2+-Mg2+-free solution (CMF, solution (c)), cells dissociate into a "sand pile" and are ready for plating. Immediately after plating into culture medium cells exhibit little morphological variability (Figure 5B) but take on distinct shapes after twenty-four hours in culture. Muscle cells become spindle-shaped while neurons remain spherical whilst extending neurites that elaborate varicosites at synapses with muscle cells (Figure 6). Simultaneous paired patch recording from the presynaptic varicosity and postsynaptic muscle cell (Figure 7) reveals the presynaptic inward and outward currents associated with the release of neurotransmitter and the resultant excitatory postsynaptic currents.
<img src="/files/ftp_upload/50253/50253fig1.jpg" alt="Figure 1"/> 
 Figure 1. A: Breeding pair of frogs in mating bucket atop screen. B: Frogs in amplexus with fertilized eggs. 
<img src="/files/ftp_upload/50253/50253fig2.jpg" alt="Figure 2"/> 
 Figure 2. A: "suitable" stage 22 embryo. B: "unsuitable" stage 28 embryo. Scale bar represents 1 mm. 
<img src="/files/ftp_upload/50253/50253fig3.jpg" alt="Figure 3"/> 
 Figure 3. A: Stage 22 embryo before removal of vitelline membrane and jelly coat. The arrow points to the outer edge of the jelly coat. The vitelline membrane adheres closely to the embryo and is virtually invisible. B: Bare embryo. Scale bar represents 1 mm.
<img src="/files/ftp_upload/50253/50253fig4.jpg" alt="Figure 4"/> 
 Figure 4. A: Stage 22 embryo with a dashed line indicating to-be dissected portion. B: Acutely isolated dorsal portion of embryo; "d" and "v" refer to the dorsal and ventral aspects of the embryos, while "m" and "n" indicate the approximate locations of the mytomes and neural tube. C: "Sand pile" of cells after 60 min in CMF. Scale bar represents 1 mm for A, and 0.5 mm for B and C.
<img src="/files/ftp_upload/50253/50253fig5.jpg" alt="Figure 5"/> 
 Figure 5. A: 5x power view of cells in culture immediately after plating. B: Same culture at 40x. Scale bar represents 40 &mu;m for A, and 5 &mu;m for B.
<img src="/files/ftp_upload/50253/50253fig6.jpg" alt="Figure 6"/> 
 Figure 6. Neuromuscular junction in culture with identification of neuronal soma (S), presynaptic varicosity (V), and postsynaptic muscle cell (M). Scale bar: 5 &mu;m.
<img src="/files/ftp_upload/50253/50253fig7.jpg" alt="Figure 7" fo:content-width="5in" fo:src="/files/ftp_upload/50253/50253fig7highres.jpg"/> 
 Figure 7. Presynaptic (top) and postsynaptic (bottom) currents seen in response to application of 10 mV incremental voltage steps presented to the presynaptic varicosity from -30 mV to +40 mV. Voltage steps are indicated next to some of the presynaptic traces. The holding potential for both cells was -70 mV.

Watch this Scientific Journal Video about Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures at JoVE.com

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

This video demonstrates the procedures used to grow primary cultures of embryonic Xenopus nerve and muscle cells and the usefulness of this preparation for making simultaneous pre- and post-synaptic patch clamp recordings.

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

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Neuroscience

Derivation of Enriched Oligodendrocyte Cultures and Oligodendrocyte/Neuron Myelinating Co-cultures from Post-natal Murine Tissues

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications for understanding the pathogenesis of demyelinating diseases such as Multiple Sclerosis (MS). Cellular development is commonly studied with primary cell culture models. Primary cell culture facilitates the evaluation of a given cell type by providing a controlled environment, free of the extraneous variables that are present in vivo. While OL cultures derived from rats have provided a vast amount of insight into OL biology, similar efforts at establishing OL cultures from mice has been met with major obstacles. Developing methods to culture murine primary OLs is imperative in order to take advantage of the available transgenic mouse lines. 
Multiple methods for extraction of OPCs from rodent tissue have been described, ranging from neurosphere derivation, differential adhesion purification and immunopurification 1-3. While many methods offer success, most require extensive culture times and/or costly equipment/reagents. To circumvent this, purifying OPCs from murine tissue with an adaptation of the method originally described by McCarthy &amp; 
de Vellis 2 is preferred. This method involves physically separating OPCs from a mixed glial culture derived from neonatal rodent cortices. The result is a purified OPC population that can be differentiated into an OL-enriched culture. This approach is appealing due to its relatively short culture time and the unnecessary requirement for growth factors or immunopanning antibodies. 
While exploring the mechanisms of OL development in a purified culture is informative, it does not provide the most physiologically relevant environment for assessing myelin sheath formation. Co-culturing OLs with neurons would lend insight into the molecular underpinnings regulating OL-mediated myelination of axons. For many OL/neuron co-culture studies, dorsal root ganglion neurons (DRGNs) have proven to be the neuron type of choice. They are ideal for co-culture with OLs due to their ease of extraction, minimal amount of contaminating cells, and formation of dense neurite beds. While studies using rat/mouse myelinating xenocultures have been published 4-6, a method for the derivation of such OL/DRGN myelinating co-cultures from post-natal murine tissue has not been described. Here we present detailed methods on how to effectively produce such cultures, along with examples of expected results. These methods are useful for addressing questions relevant to OL development/myelinating function, and are useful tools in the field of neuroscience.

This article describes methods to derive enriched populations of murine oligodendrocyte precursor cells (OPCs) in primary culture, which differentiate to produce mature oligodendrocytes (OLs). In addition, this report describes techniques to produce murine myelinating co-cultures by seeding mouse OPCs onto a neurite bed of mouse dorsal root ganglion neurons (DRGNs).

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

Forebrain Electrophysiological Recording in Larval Zebrafish

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

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

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

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

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

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

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

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

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

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

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

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

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

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

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

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the researcher to measure gustatory responses directly and quantitatively, reflecting the sensory input that the insect nervous system receives from taste stimuli in its environment. This protocol outlines all key steps in performing this technique. The critical steps in assembling an electrophysiology rig, such as selection of necessary equipment and a suitable environment for recording, are delineated. We also describe how to prepare for recording by making appropriate reference and recording electrodes, and tastant solutions. We describe in detail the method used for preparing the insect by insertion of a glass reference electrode into the fly in order to immobilize the proboscis. We show traces of the electrical impulses fired by taste neurons in response to a sugar and a bitter compound. Aspects of the protocol are technically challenging and we include an extensive description of some common technical challenges that may be encountered, such as lack of signal or excessive noise in the system, and potential solutions. The technique has limitations, such as the inability to deliver temporally complex stimuli, observe background firing immediately prior to stimulus delivery, or use water-insoluble taste compounds conveniently. Despite these limitations, this technique (including minor variations referenced in the protocol) is a standard, broadly accepted procedure for recording Drosophila neuronal responses to taste compounds.

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

This protocol describes extracellular recording of the action potential responses fired by labellar taste neurons in Drosophila.

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the electrical responses of the retina to light stimulation, while the VEP measures the corresponding functional integrity of the visual pathways from the retina to the primary visual cortex following the same light event. The ERG waveform can be broken down into components that reflect responses from different retinal neuronal and glial cell classes. The early components of the VEP waveform represent the integrity of the optic nerve and higher cortical centers. These recordings can be conducted in isolation or together, depending on the application. The methodology described in this paper allows simultaneous assessment of retinal and cortical visual evoked electrophysiology from both eyes and both hemispheres. This is a useful way to more comprehensively assess retinal function and the upstream effects that changes in retinal function can have on visual evoked cortical function.

This protocol describes simultaneous measurement of electroretinogram and visual evoked potentials in anesthetized rats.

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the ...

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, oviposition, and mating. Myriad chemicals of varying volatilities have been identified as natural odorants that activate insect Olfactory Receptor Neurons (ORNs). However, studying the olfactory responses to low-volatility odorants has been hampered by an inability to effectively present such stimuli using conventional odor-delivery methods. Here, we describe a procedure that permits the effective presentation of low-volatility odorants for in vivo Single-Sensillum Recording (SSR). By minimizing the distance between the odor source and the target tissue, this method allows for the application of biologically salient but hitherto inaccessible odorants, including palmitoleic acid, a stimulatory pheromone with a demonstrated effect on ORNs involved in courtship and mating behavior1. Our procedure thus affords a new avenue to assay a host of low-volatility odorants for the study of insect olfaction and pheromone communication.

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

The overall goal of this protocol is to demonstrate how to present odorants of low volatility for single-sensillum recording from Drosophila olfactory receptor neurons that respond to long-chain cuticular pheromones.

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, ...