We thank Dr. Meng-Chin Lin and Ms. Yuan Dong for technical assistance. This work was supported by a grant from the National Institutes of Health HD053767 (subcontract to NLW), and by funds from the Department of Physiology and Office of the Vice Chancellor for Research, University of California-Los Angeles (NLW).

1. Dissection of Brains from Adult Medaka
<ol>
	<li>Anesthetize adult male or female (Figure 1A) by immersing in 5 ml MS-222 (150 mg/L, pH 7.4); wait a couple of minutes after gills movements have ceased before decapitating. All procedures were approved by the Institutional Animal Care and Use Committee of University of California-Los Angeles.</li>
	<li>Decapitate the fish in fish saline at the caudal end of the operculum with scissors in a 60-mm diameter Petri dish.</li>
	<li>Transfer fish head to a 35-mm diameter Petri dish, half filled with fish saline and lined with Sylgard that has a depression in it to position and secure the head for brain dissection. Position and secure the head dorsal-side up with insect pins.</li>
	<li>Carefully cut the dorsal part of the skull around its perimeter with fine scissors and gently remove it with forceps, exposing the dorsal part of brain completely including the anterior part of the spinal cord.</li>
	<li>Lift the spinal cord gently with fine-tip forceps and sever the bilateral connective nerves with fine-tipped scissors: cranial nerves, spinal nerves, and optic nerves on the ventral side of the brain, and the olfactory nerves at the anterior part of the brain.</li>
	<li>Place the fully dissected brain in a fresh Petri dish filled with fish saline, and check carefully under the microscope to ensure the whole brain is intact (including the pituitary gland) without any cuts or punctures. Damaged brains are not used for experimentation.</li>
</ol>
2. Mounting the Brain in a Recording Chamber
<ol>
	<li>Place a small drop of cyanoacrylate or some other fast-acting glue with a needle into the center of the recording chamber and spread the glue about 1 cm2.</li>
	<li>Quickly transfer the brain and glue it down in a ventral-side up position.</li>
	<li>Add 1 ml of fish saline so that it fills the recording chamber and covers the brain. Continuously perfuse the brain with aerated fish saline at a rate of at least 200 &mu;l/min.</li>
	<li>Using the fine-tip forceps carefully remove the meninges over the surface of the recording area of the brain.</li>
</ol>
3. Electrophysiology
<ol>
	<li>Loose patch recording of action potential firing
	<ol>
		<li>Glass microelectrodes (6-10 M&Omega;) are constructed from borosilicate glass pipettes (1B150F-4, World Precision Instruments) using an electrode puller (e.g. Sutter P-87), and back filled with the loose-patch recording solution half-way up the electrode.</li>
		<li>Using an upright fluorescent microscope with a cooled charge-coupled device (CCD) camera, find the GFP-expressing TN-GnRH3 neurons through 10x (Figure 1B), then 40x water immersion objectives (Figure 1C). They are located at or near the ventral surface of the olfactory bulbs, just anterior to the telencephalon.</li>
		<li>Switch to a video monitor that is providing real-time images from the microscope to locate the TN-GnRH3 neuron cluster (Figure 1C).</li>
		<li>Position the microelectrode into the bath so that the tip of the electrode is above the neuron target. Apply constant slight positive pressure to the microelectrode so that it doesn't become clogged, and gently approach the target neuron with the tip of the electrode.</li>
		<li>Check and monitor the electrode resistance in voltage-clamp mode using AxoGraph software and amplifier Axoclamp 200B with Axograph software (Axon Instruments). When you can see from the video monitor that the tip of the electrode is on the surface of the neuron and from the AxoGraph display that the resistance changes slightly, release the positive pressure and apply slight negative pressure if necessary to make a low resistance seal on the neuron (&lt;100 M&Omega;).</li>
		<li>Switch to current-clamp mode, and record the membrane voltage continuously without any current input, adjusting the scale of voltage and rate of recording if necessary. Data is collected and analyzed by Powerlab software (ADInstruments Inc.).</li>
		<li>Record baseline electrical activity in normal fish saline for about 5 min before any treatment (Figure 2A). Bath perfusion of drugs, hormones, peptides, etc. added to fish saline continues at the rate of 200 &mu;l/min, followed by a washout period in normal fish saline 5, 6, 11.</li>
	</ol>
	</li>
	<li>Whole cell recording of membrane potential 
	The whole cell patch clamp electrophysiology procedure is the same as that of the extracellular loose-patch recording procedure described above, from steps 3.1.1 - 3.1.4. Then the procedures deviate accordingly:
	<ol>
		<li>Check and monitor the electrode resistance in voltage-clamp mode. When you can see from the video monitor that the tip of the electrode is on the surface of the neuron and from the oscilloscope that the resistance changes slightly, release the positive pressure and apply a little negative pressure to make a high resistance seal on the neuron (~ 3 Giga &Omega;). Apply gentle suction by mouth to rupture the patched membrane; you will see a sudden drop of the resistance to about 120-250 M&Omega;.</li>
		<li>Switch to current-clamp mode, and record the membrane voltage continuously without any current input, adjusting the scale of voltage and rate of recording if necessary.</li>
		<li>Record baseline electrical activity in normal fish saline for about 5 min before any treatment, as described above in 3.1.7 (Figure 2B).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Kah, O., Lethimonier, C., Lareyre, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gonadotrophin-releasing+hormone+(GnRH)+in+the+animal+kingdom.">Gonadotrophin-releasing hormone (GnRH) in the animal kingdom.</a> J. Soc. Biol. 198 (1), 53-60 (2004).</li><li>Ramakrishnan, S., Wayne, N. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Social+cues+from+conspecifics+alter+electrical+activity+of+gonadotropin-releasing+hormone+neurons+in+the+terminal+nerve+via+visual+signals.">Social cues from conspecifics alter electrical activity of gonadotropin-releasing hormone neurons in the terminal nerve via visual signals.</a> Am. J. Physiol. Regul. Integr. Comp. Physiol. 297 (1), R135-R141 (2009).</li><li>Abe, H., Oka, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+neuromodulation+by+a+nonhypophysiotropic+GnRH+system+controlling+motivation+of+reproductive+behavior+in+the+teleost.">Mechanisms of neuromodulation by a nonhypophysiotropic GnRH system controlling motivation of reproductive behavior in the teleost.</a> 57 (6), 665-674 (2011).</li><li>Oka, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tetrodotoxin-resistant+persistent+Na++current+underlying+pacemaker+potentials+of+fish+gonadotrophin-releasing+hormone+neurones.">Tetrodotoxin-resistant persistent Na+ current underlying pacemaker potentials of fish gonadotrophin-releasing hormone neurones.</a> J. Physiol. 482 (Pt. 1), 1-6 (1995).</li><li>Zhao, Y., Wayne, N. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+Kisspeptin1+on+Electrical+Activity+of+an+Extrahypothalamic+Population+of+Gonadotropin-Releasing+Hormone+Neurons+in+Medaka.">Effects of Kisspeptin1 on Electrical Activity of an Extrahypothalamic Population of Gonadotropin-Releasing Hormone Neurons in Medaka.</a> PLoS One. 7 (5), e37909 (2012).</li><li>Wayne, N. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-cell+electrophysiology+of+gonadotropin-releasing+hormone+neurons+that+express+green+fluorescent+protein+in+the+terminal+nerve+of+transgenic+medaka+(Oryzias+latipes).">Whole-cell electrophysiology of gonadotropin-releasing hormone neurons that express green fluorescent protein in the terminal nerve of transgenic medaka (Oryzias latipes).</a> Biol. Reprod. 73 (6), 1228-1234 (2005).</li><li>Okubo, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Forebrain+gonadotropin-releasing+hormone+neuronal+development:+insights+from+transgenic+medaka+and+the+relevance+to+X-linked+Kallmann+syndrome.">Forebrain gonadotropin-releasing hormone neuronal development: insights from transgenic medaka and the relevance to X-linked Kallmann syndrome.</a> Endocrinology. 147 (3), 1076-1084 (2006).</li><li>Okubo, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+form+of+gonadotropin-releasing+hormone+in+the+medaka,+Oryzias+latipes.">A novel form of gonadotropin-releasing hormone in the medaka, Oryzias latipes.</a> Biochem. Biophys. Res. Commun. 276 (1), 298-303 (2000).</li><li>Ramakrishnan, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acquisition+of+spontaneous+electrical+activity+during+embryonic+development+of+gonadotropin-releasing+hormone-3+neurons+located+in+the+terminal+nerve+of+transgenic+zebrafish+(Danio+rerio).">Acquisition of spontaneous electrical activity during embryonic development of gonadotropin-releasing hormone-3 neurons located in the terminal nerve of transgenic zebrafish (Danio rerio).</a> Gen. Comp. Endocrinol. 168 (3), 401-407 (2010).</li><li>Abraham, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+gonadotropin-releasing+hormone-3+neuron+ablation+in+zebrafish:+effects+on+neurogenesis,+neuronal+migration,+and+reproduction.">Targeted gonadotropin-releasing hormone-3 neuron ablation in zebrafish: effects on neurogenesis, neuronal migration, and reproduction.</a> Endocrinology. 151 (1), 332-340 (2010).</li><li>Wayne, N. L., Kuwahara, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Beta-endorphin+alters+electrical+activity+of+gonadotropin+releasing+hormone+neurons+located+in+the+terminal+nerve+of+the+teleost+medaka+(Oryzias+latipes.">Beta-endorphin alters electrical activity of gonadotropin releasing hormone neurons located in the terminal nerve of the teleost medaka (Oryzias latipes.</a> Gen. Comp. Endocrinol. 150 (1), 41-47 (2007).</li><li>Oka, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three+types+of+gonadotrophin-releasing+hormone+neurones+and+steroid-sensitive+sexually+dimorphic+kisspeptin+neurones+in+teleosts.">Three types of gonadotrophin-releasing hormone neurones and steroid-sensitive sexually dimorphic kisspeptin neurones in teleosts.</a> J. Neuroendocrinol. 21 (4), 334-338 (2009).</li><li>Molleman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Patch Clamping: An Introductory Guide To Patch Clamp Electrophysiology. , (2003).</li></ol>

The authors have nothing to disclose.

GnRH3:GFP transgenic fishes provide unique models to study the neurophysiological mechanisms underlying neuronal integration and regulation in central control of behaviors that are both directly and indirectly involved in reproduction3, 8-10. One of the significant advantages of this model system is that many GnRH3 neurons expressing GFP are close to the ventral surface of the brain, allowing for relatively easy access to the neurons for electrophysiological recording without disrupting neural circuits6, 9, 11, 12 (Figures 1B and 1C; Figures 3A and 3B). In this video, we have shown both loose-patch and whole-cell recordings from TN-GnRH3 neurons in the intact adult brain of medaka fish. TN-GnRH3 neurons exhibit a tonic or beating pattern of spontaneous action potential firing with a frequency of 0.5-6Hz.There were no detectable sex differences in spontaneous firing rate 5, but there were dramatic developmental changes in GnRH3 neuron electrical activities during embryogenesis9. Loose-patch and whole-cell electrophysiology have different methodological advantages and disadvantages. Loose-patch is technically easier than whole-cell recording, but only shows the neuron firing pattern and frequency. On the other hand, whole-cell recording can provide much more information: resting membrane potential, firing pattern and frequency, action potential analysis, membrane capacitance and resistance. Because loose-patch recording does not break the cell membrane and disrupt the intracellular ion and second messenger concentrations, it is ideal for long-term recording of neuronal activity. Whole cell recordings rupture the cell membrane and eventually lead to dialysis of intracellular molecules, thereby potentially altering neuron properties with more prolonged recordings. Because of the advantages and disadvantages of both electrophysiological methods, it can be important to study neuron properties using both techniques. Because both pH and osmolarity are very important for neuron survival and function, all the solutions in use need to be adjusted properly.
To confirm the identity of the recorded neuron, negative pressure is applied after completion of the recording, and then fluorescence at the tip of the electrode is confirmed using microscopic fluorescence imaging. Small molecular dyes or probes (such as Alexa Fluor 568 dye or biocytin) may be included in the intracellular solution for the whole-cell recording to mark the recorded neuron for further morphological analysis.
To record successfully from target neurons in intact brain, the depth of location of the neurons is crucial -- the deeper the neurons, the harder to patch (both loose-patch and whole-cell). Generally, the feasible depth for successful patching is about 5-6 cell layers from the surface. Medaka TN-GnRH3 neurons are typically at the ventral surface of the brain, while zebrafish hypothalamic GnRH3 neurons can be up to 6 cell layers from the surface. Positive pressure is very helpful to prevent the recording electrode from clogging while approaching the cells. This type of experimental approach can be used for any neuron that is genetically labeled with fluorescent protein. To study GnRH neuron activity in the adult brain, we routinely use medaka fish and zebrafish of 4-6 months of age. The younger fish with smaller brains or older fish with more calcified skulls, will make the dissection more challenging. The TN:GnRH3 neurons are easily accessible for electrophysiology from adult medaka brain, but this is not the case for adult zebrafish in which TN:GnRH3 neurons are encased in a tough connective sheath. On the other hand, GnRH3 neurons in POA, VT, and hypothalamus are more readily accessible for electrophysiology in adult zebrafish brain than from medaka. There are no sex differences based on the difficulties of dissection and electrophysiology recording of either species.
Although this is not an in vivo experimental approach, recording from the relatively intact neuronal circuits with minimum damage is an attractive way to explore the physiological function and regulation of the neuronal system of interest. The intact brain preparation lasts for hours, and a good recording can last for over an hour 6. This protocol is suitable for researchers who already have basic knowledge of and experience with electrophysiology techniques 13, and as well as the specialized equipment required to do the work.

<table border="1">
 <tbody>
 <tr>
 <td>Name</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Microscope</td>
 <td>Olympus</td>
 <td></td>
 <td>BX50W (Upright)</td>
 </tr>
 <tr>
 <td>Amplifier</td>
 <td>Axon Instruments</td>
 <td></td>
 <td>Axoclamp 200B </td>
 </tr>
 <tr>
 <td>A-D converter</td>
 <td>Computer Interference Corp.</td>
 <td></td>
 <td>Digidata ITC-18 </td>
 </tr>
 <tr>
 <td>Cooled CCD camera</td>
 <td>PCO Computer Optics </td>
 <td></td>
 <td>Sensicam</td>
 </tr>
 <tr>
 <td>Xenon lamp</td>
 <td>Sutter Instruments Co.</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>GFP filter set</td>
 <td>Chroma Technologies</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Imaging Software</td>
 <td>Intelligent Imaging Innovations</td>
 <td></td>
 <td>Slidebook software </td>
 </tr>
 <tr>
 <td>Electrophysiology Data Acquisition Software</td>
 <td>Axon Instruments</td>
 <td></td>
 <td>Axograph software </td>
 </tr>
 <tr>
 <td>Electrophysiology Data Acquisition Software</td>
 <td>AD Instruments Inc.</td>
 <td></td>
 <td>PowerLab </td>
 </tr>
 <tr>
 <td>Headstage for electrophysiology</td>
 <td>Axon Instruments</td>
 <td></td>
 <td>CV 203BU </td>
 </tr>
 <tr>
 <td>Micromanipulator</td>
 <td>Sutter Instrument Co </td>
 <td>MP-285 </td>
 <td></td>
 </tr>
 <tr>
 <td>Recording Chamber Platform</td>
 <td>Warner Instrument Corp.</td>
 <td></td>
 <td>P1 </td>
 </tr>
 <tr>
 <td>Recording Chamber</td>
 <td>Warner Instrument Corp.</td>
 <td></td>
 <td>RC-26G </td>
 </tr>
 <tr>
 <td>Electrode Puller</td>
 <td>Sutter instruments</td>
 <td></td>
 <td>P87 </td>
 </tr>
 <tr>
 <td>Filament for electrode puller</td>
 <td>Sutter Instruments</td>
 <td>FB330B</td>
 <td>3.0 mm wide trough filament</td>
 </tr>
 <tr>
 <td>1.5 mm glass capillaries</td>
 <td>World Precision Instruments</td>
 <td>1B150-4</td>
 <td>Microelectrode for recording</td>
 </tr>
 <tr>
 <td>Syringe</td>
 <td>Becton Dickinson</td>
 <td>309586</td>
 <td>3 ml</td>
 </tr>
 <tr>
 <td>MS-222 </td>
 <td>Sigma</td>
 <td>E10521-10G</td>
 <td>Ethyl 3-aminobenzoate methanesulfonate salt</td>
 </tr>
 <tr>
 <td>Fish saline</td>
 <td></td>
 <td></td>
 <td>mM: 134 NaCl; 2.9 KCl; 2.1 CaCl2; 1.2 MgCl2; 10 HEPES </td>
 </tr>
 <tr>
 <td>Electrode solution (loose-patch) </td>
 <td></td>
 <td></td>
 <td>mM: 150 NaCl; 3.5 KCl; 2.5 CaCl2; 1.3 MgCl2; 10 HEPES; 10 glucose </td>
 </tr>
 <tr>
 <td>Electrode solution (whole-cell patch) </td>
 <td></td>
 <td></td>
 <td>mM: 112.5 K-gluconate; NaCl; 17.5 KCl; 0.5 CaCl2; 1 MgCl2; 5 MgATP; 1 EGTA; 10 HEPES; 1 GTP; 0.1 leupeptin;10 phospho-creatine </td>
 </tr>
 </tbody>
</table>

recording electrical activity from identified neurons in the intact brain of transgenic fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

An example of bilateral clusters of GFP-labeled TN-GnRH3 neurons from the excised brain of medaka fish are shown in Figures 1B and 1C. Each cluster contains about 8-10 GnRH neurons. The spontaneous neuronal activities of the target TN-GnRH3 were recorded in current-clamp mode (I=0) with typical firing rates of 0.5-6 Hz. The pattern of action potential firing is typically a tonic or beating pattern, with a fairly regular interspike interval. Sample traces are shown in Figure 2 (2A: loose-patch; 2B: whole-cell).
This experimental approach is also successful in recording from GFP-labeled GnRH neurons located in the forebrain of adult zebrafish (Figures 3A and 3B). Neuron electrical activity in the excised, intact brain was recorded in the similar way as described above in medaka by loose-patch (Figure 3C) and whole-cell recording (Figure 3D). Unlike with medaka TN-GnRH neurons, the pattern of spontaneous action potential firing of zebrafish GnRH neurons in the preoptic area, ventral telencephalon, and hypothalamus is often irregular.
<img alt="Figure 1" src="/files/ftp_upload/50312/50312fig1.jpg" /> 
 Figure 1. Animal model used for the TN-GnRH3 neuron study in this video. A: Adult medaka GnRH3:GFP transgenic fish, left: male; right: female. B: Confocal image of the ventral view of the forebrain. OB: olfactory bulb; TN: Terminal nerve GnRH3 neurons; TELE: telencephalon; ON: optic nerve. C: High magnification of TN-GnRH3 neurons shown in white box of panel B. (Scale bar: A: 5 mm; B: 100 &mu;m; C: 20 &mu;m).
<img alt="Figure 2" src="/files/ftp_upload/50312/50312fig2.jpg" /> 
 Figure 2. Spontaneous action potentials recorded from TN-GnRH3 neurons in current-clamp mode (I=0). A: Sample trace from loose-patch recording; B: Sample trace from whole-cell recording.
<img alt="Figure 3" fo:content-width="6in" fo:src="/files/ftp_upload/50312/50312fig3highres.jpg" src="/files/ftp_upload/50312/50312fig3.jpg" /> 
 Figure 3. Recording of GnRH3:EMD (Emerald green fluorescent protein) neuron activity from intact brain preparation of adult transgenic zebrafish9 using the same experimental approach as described for the medaka brain preparation. A: GnRH3:EMD neurons located in ventral telencephalon (VT) and preoptic area (POA) in forebrain of adult zebrafish. B: GnRH3:EMD neurons located in hypothalamus (HYPO). Scale bars: 100 &mu;m. C: Sample trace of loose-patch recording from GnRH3:EMD neuron in VT. D: Sample trace of whole-cell patch recording from VT.

Watch this Scientific Journal Video about Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish at JoVE.com

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

recording-electrical-activity-from-identified-neurons-intact-brain

Nanyang Technological University

Research

JoVE Journal

Neuroscience

13.4K Views.  University of California, Los Angeles. In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Video: Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

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, ...

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 ...

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of neurophysiology, neuroethology, and developmental biology. The goal of this report is to consolidate experimental techniques from the leech system into a single article that will be of use to physiologists with expertise in other nervous system preparations, or to biology students with little or no electrophysiology experience. We demonstrate how to dissect the leech for recording intracellularly from identified neural circuits in the ganglion. Next we show how individual cells of known function can be removed from the ganglion to be cultured in a Petri dish, and how to record from those neurons in culture. Then we demonstrate how to prepare a patch of innervated skin to be used for mapping sensory or motor fields. These leech preparations are still widely used to address basic electrical properties of neural networks, behavior, synaptogenesis, and development. They are also an appropriate training module for neuroscience or physiology teaching laboratories.

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

This article describes three nervous system preparations using leeches: intracellular recording from neurons in ventral ganglia, culturing neurons from ventral ganglia, and recording from a patch of innervated skin to map sensory fields.

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of ...

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 Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant Receptor

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of olfactory-driven behaviors and their modulation. These coding properties depend heavily on the initial interaction between odor molecules and the olfactory receptor (OR) expressed in the OSNs. The identity, specificity and ligand spectrum of the expressed OR are critical. The probability to find the ligand of the OR expressed in an OSN chosen randomly within the epithelium is very low. To address this challenge, this protocol uses genetically tagged mice expressing the fluorescent protein GFP under the control of the promoter of defined ORs. OSNs are located in a tight and organized epithelium lining the nasal cavity, with neighboring cells influencing their maturation and function. Here we describe a method to isolate an intact olfactory epithelium and record through patch-clamp recordings the properties of OSNs expressing defined odorant receptors. The protocol allows one to characterize OSN membrane properties while keeping the influence of the neighboring tissue. Analysis of patch-clamp results yields&#160;a precise quantification of ligand/OR interactions, transduction pathways and pharmacology, OSNs&#39; coding properties and their modulation at the membrane level.&#160;

Analyzing the physiological properties of olfactory sensory neurons still faces technical limitations. Here we record them through perforated patch-clamp in an intact preparation of the olfactory epithelium in gene-targeted mice. This technique allows the characterization of membrane properties and responses to specific ligands of neurons expressing defined olfactory receptors.

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

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

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

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

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

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

Recording Brain Electromagnetic Activity During the Administration of the Gaseous Anesthetic Agents Xenon and Nitrous Oxide in Healthy Volunteers

Anesthesia arguably provides one of the only systematic ways to study the neural correlates of global consciousness/unconsciousness. However to date most neuroimaging or neurophysiological investigations in humans have been confined to the study of &#947;-Amino-Butyric-Acid-(GABA)-receptor-agonist-based anesthetics, while the effects of dissociative N-Methyl-D-Aspartate-(NMDA)-receptor-antagonist-based anesthetics ketamine, nitrous oxide (N2O) and xenon (Xe) are largely unknown. This paper describes the methods underlying the simultaneous recording of magnetoencephalography (MEG) and electroencephalography (EEG) from healthy males during inhalation of the gaseous anesthetic agents N2O and Xe. Combining MEG and EEG data enables the assessment of electromagnetic brain activity during anesthesia at high temporal, and moderate spatial, resolution. Here we describe a detailed protocol, refined over multiple recording sessions, that includes subject recruitment, anesthesia equipment setup in the MEG scanner room, data collection and basic data analysis. In this protocol each participant is exposed to varying levels of Xe and N2O in a repeated measures cross-over design. Following relevant baseline recordings participants are exposed to step-wise increasing inspired concentrations of Xe and N2O of 8, 16, 24 and 42%, and 16, 32 and 47% respectively, during which their level of responsiveness is tracked with an auditory continuous performance task (aCPT). Results are presented for a number of recordings to highlight the sensor-level properties of the raw data, the spectral topography, the minimization of head movements, and the unequivocal level dependent effects on the auditory evoked responses. This paradigm describes a general approach to the recording of electromagnetic signals associated with the action of different kinds of gaseous anesthetics, which can be readily adapted to be used with volatile and intravenous anesthetic agents. It is expected that the method outlined can contribute to the understanding of the macro-scale mechanisms of anesthesia by enabling methodological extensions involving source space imaging and functional network analysis.

Simultaneous magnetoencephalography and electroencephalography provides a useful tool to search for common and distinct macro-scale mechanisms of reductions in consciousness induced by different anesthetics. This paper illustrates the empirical methods underlying the recording of such data from healthy humans during N-Methyl-D-Aspartate-(NMDA)-receptor-antagonist-based anesthesia during inhalation of nitrous oxide and xenon.

Anesthesia arguably provides one of the only systematic ways to study the neural correlates of global consciousness/unconsciousness. However to date most ...