I thank L Watanabe, M Gorczyca and other members of the Yoshihara lab for useful comments and discussion. I thank K. Scott and L. Looger for fly stocks, S Yokoyama for demonstrating experiments, Shinya Iguchi for technical help, and Nobuko Yoshihara for material information. This work was supported by National Institute of Mental Health Grants MH85958, and the Worcester Foundation to M.Y.

1. Constructing the FLIES
<ol>
	<li>Shape a platform from the lid of 35 mm Falcon dish by melting the sidewall and carving it to form an appropriate angle and thickness (Figure 1a; Figure 2). Drill a hole in the platform to accept the fly head (Figure 1a,b), such that the mouthparts are freely exposed to the outside of the chamber (Figure 1a). Cut the end of a Pipetman tip, with the size matching the fly to be observed. Glue the tip to the platform as shown in Figure 1a to complete the FLIES (Figure 2).</li>
</ol>
2. Preparing the Fly for Observation
<ol>
	<li>Starve an adult fly for 24 hours at 25 &deg;C prior to the experiment by placing it in a vial with only a wet paper towel, if starvation is necessary.</li>
	<li>Anesthetize the fly by placing it in a 15ml plastic tube standing on ice.</li>
	<li>Using forceps, insert a fly into the chamber of the FLIES, and gently push the fly in until it is unable to move. Then, insert a plug to retain the fly. (Figure 1a,b).</li>
	<li>Seal the surrounding parts of the proximal proboscis (rostrum) to the inner edge of the hole; Apply a small amount of light-curing glue (Tetric EvoFlow) to the sides and above the rostrum from the outside (arrows in Figure 3). Also apply the glue from the inside of the FLIES to the space between the dorsal part of the head and the inner edge of the hole (Figure 1b). To allow the rostrum to move freely, care should be taken to prevent any glue from touching the rostrum by using an eyelash (or something comparable) to spread the glue. Finally, cure the glue with the weakest illumination of blue light sufficient for curing to avoid damaging the fly from excessive heat. Remove the plug.</li>
	<li>A thread to hold the rostrum partially lifted (arrow in Figure 3) may be employed to stabilize the fly's head by avoiding bump of the pharyngeal part to SOG and to expose the ventral part of the SOG, especially for imaging presynaptic terminals of Gr5a neurons8, which are covered by part of the pharynx when the proboscis is fully retracted.</li>
	<li>Fill the surface of the platform with a sugar-free saline containing (in mM): NaCl, 140; KCl, 2; MgCl2, 4.5; CaCl2, 1.5; and HEPES-NaOH, 5 with a pH of 7.110. This sucrose-free saline has similar tonicity to those of commonly-used sucrose-containing salines such as HL3.111.</li>
	<li>Open the head capsule using a tungsten blade and forceps with sharpened tips like scissors, which is a custom-made device and originally designed by Dr. Kageyuki Yamaoka, Japan, to clip the cuticle and trachea to expose the SOG (Figure 4). The ventral edge of the head cuticle was cut as ventral as possible whereas the dorsal edge cut was not as critical. First, make an incision through the posterior edge using a tungsten blade. Then, cut through the side and the anterior edges using the sharpened forceps. The cut cuticle piece should be lifted with the forceps and removed. Antennae and antennal nerves should be cut out by the sharpened forceps.</li>
	<li>In order to avoid movement artifacts during recordings, the brain must be stabilized through the removal of certain parts of the fly. First, selectively remove air sacks between the SOG and cuticle. Cut the oesophagus at the end toward the pharynx and at the end going into the SOG to remove the connection between the pharynx and the brain. Then pull out Muscle 1612, and finally, detach any trachea connecting the brain and the cuticle.</li>
	<li>We set our FLIES on the stage of an Olympus BX51WI microscope connected to a spinning disk confocal microscope system (Improvision in PerkinElmer) (Figure 5). A water immersion lens, e.g. 40X/0.8 N.A, was immersed into the bath (Figure 6).</li>
</ol>
3. Ca2+ Imaging of the Brain
<ol>
	<li>Ca2+ imaging using GCaMP3.01, 9 was performed through the method established by Marella et al.8 (Figure 6). Using a spinning disk confocal microscope (Improvision), take a single optical section at 4Hz with an exposure time of 122ms using 491nm excitation laser, focusing at the region of interest (a cell body of a motor neuron, or an interneuron, or presynaptic terminals of gustatory sensory neurons) with Velocity software, ver. 4.3 (Improvision).</li>
</ol>
4. Stimulating the Proboscis
<ol>
	<li>Aspirate 100mM aqueous sucrose solution into a hypodermic needle with a small wick (made from Japanese Washi paper, see Table of specific reagents) protruding from the tip (Figure 7).</li>
	<li>Discharge a small drop of sucrose solution onto the wick, then, aspirate it, immediately before applying the soaked Washi paper wick to the tip of the proboscis. The Washi paper wick should only be applied for an instant to avoid satiation (Figure 8). Monitor the Proboscis Extension Response (PER) behavior using a CCD camera attached to a dissection microscope (Figure 5) simultaneously with Ca2+ imaging. Data must be taken within one hour after starting the dissection because the PER response is not maintained more than one hour under these conditions.</li>
</ol>
5. Representative Results
Motor neurons of the rostrum protractor, a pair of major muscles responsible for lifting the rostrum for proboscis extension, have been reported to show robust responses to sweet stimuli13. Figure 9 shows Ca2+ signals through GCaMP3.01, 9 expressed by a Gal 4 driver, E4913, from a cell body of the motor neuron. As shown in the video, a robust PER was observed in synchrony with an increase in Ca2+ signal.
<img alt="Figure 1" src="/files/ftp_upload/3625/3625fig1.jpg" /> 
Figure 1. Schematic of the Fly brain Live Imaging and Electrophysiology Stage (FLIES). a, The FLIES is built such that the angle of the wall allows for the fly head to come out straight. Three segments of the fly proboscis are color-coded; the rostrum is magenta. The fly is inserted into the chamber with forceps, and a piece cut from the end of a Pipetman tip is plugged into the back of the chamber to block the fly from escaping. b, The hole is bored to conform to the fly's head. The remaining gaps are sealed with light-curing glue as indicated to ensure that there are no leaks. After the glue has cured, the Pipetman tip as a plug is removed. Saline is poured into the FLIES such that the surface of the brain is completely covered. It should be ensured that there is no saline leaking out onto the anterior end of the fly head. The shaded area surrounded by the dashed line in panel b shows the part to be dissected out.
<img alt="Figure 2" src="/files/ftp_upload/3625/3625fig2.jpg" /> 
Figure 2. Photograph of the FLIES. The crescent shaped wall of glue (from a glue gun) is made to control the flow of saline solution during perfusion, and to reduce the amount of saline used.
<img alt="Figure 3" src="/files/ftp_upload/3625/3625fig3.jpg" /> 
Figure 3. Frontal view of a fly mounted in the FLIES. This image shows how a fly is set in the FLIES, and how the light-curing glue is applied around the head of the fly (arrows) to create a saline-proof seal. If any saline leaks through to the proboscis of the fly, then the experiment will fail. The thread (arrowheads) tethered to the stage is to hold the proboscis in position, so that it is not completely retracted. Otherwise it would obstruct the more ventral portion of the SOG and cause movement artifacts.
<img alt="Figure 4" src="/files/ftp_upload/3625/3625fig4.jpg" /> 
Figure 4. Top-view of the FLIES with a dissected fly. This image shows the exposed SOG of a fly following cuticle removal. The esophagus, Muscle 16 and tracheae connected to the brain, as well as selected air-sacks have also been removed to prevent them from disturbing the brain, which can lead to artifacts in the calcium signals.
<img alt="Figure 5" src="/files/ftp_upload/3625/3625fig5.jpg" /> 
Figure 5. The monitoring apparatus. This apparatus is assembled from an Olympus BX51WI microscope attached to a spinning disk confocal microscope system (Improvision), and a side-mounted Nikon SMZ800 microscope. This setup allows for the live-imaging of brain activity during PER behavior.
<img alt="Figure 6" src="/files/ftp_upload/3625/3625fig6.jpg" /> 
Figure 6. Photographs to show the side-view of the FLIES. The thin layer of saline solution is sandwiched between the 40X water-immersion lens and the fly.
<img alt="Figure 7" src="/files/ftp_upload/3625/3625fig7.jpg" /> 
Figure 7. Schematic for making the Washi paper wick. Gampi-Washi paper is a traditional Japanese parchment that is extremely thin and smooth (Haibara, Japan). It is also very strong and can retain a large amount of liquid. The Washi paper must be cut into a trapezoid with very specific dimensions; 0.3 mm and 0.7 mm in width, and 4-5 mm in length (a). This trapezoid is then inserted from the 0.7 mm side into the tip of a 23 G hypodermic needle and can also be stabilized through the insertion of an insect pin, which is bent to a V-shape (b). c, The Washi paper becomes transparent following exposure to a liquid, making it ideal for this experiment.
<img alt="Figure 8" src="/files/ftp_upload/3625/3625fig8.jpg" /> 
Figure 8. Stimulation of the proboscis on the FLIES with the Washi-wick. The Washi paper wick at the tip of a hypodermic needle is applied for an instant using a Joystick manipulator with one hand. The needle is connected through a flexible tube to an injector manipulated with the other hand, for aspirating and discharging sucrose solution.
<img alt="Figure 9" src="/files/ftp_upload/3625/3625fig9.jpg" /> 
Figure 9. Representative results. a, PER behavior recorded through the CCD camera installed on the stereomicroscope. Frontal views of a starved fly with an extended proboscis (arrows) before the stimulation, at the stimulation, and after the stimulation with a wick containing 100 mM sucrose aqueous solution (arrowhead). Illumination is accomplished by a 491 nm laser used for GCaMP fluorescence. b, Sucrose-induced GCaMP 3.0 response in the motor neuron for the protractor of rostrum muscle in the starved state. The top panel, before the stimulation; the bottom panel, immediately after the stimulation. Scale bar, 10 &mu;m. c, Quantification of the sucrose-induced GCaMP 3.0 response. The motor neuron responds to a sucrose stimulus by a sharp increase in fluorescence followed by a restoration phase of several seconds to the base line.

<ol>
	<li>Nakai, J., Ohkura, M., Imoto, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+high+signal-to-noise+Ca(2+)+probe+composed+of+a+single+green+fluorescent+protein.">A high signal-to-noise Ca(2+) probe composed of a single green fluorescent protein.</a> Nat. Biotechnol. 19, 137-141 (2001).</li><li>Lima, S. Q., Miesenbock, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Remote+control+of+behavior+through+genetically+targeted+photostimulation+of+neurons.">Remote control of behavior through genetically targeted photostimulation of neurons.</a> Cell. 121, 141-152 (2005).</li><li>Schroll, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light-induced+activation+of+distinct+modulatory+neurons+triggers+appetitive+or+aversive+learning+in+Drosophila+larvae.">Light-induced activation of distinct modulatory neurons triggers appetitive or aversive learning in Drosophila larvae.</a> Curr. Biol. 16, 1741-1747 (2006).</li><li>Hamada, F. 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I have nothing to disclose.

The FLIES allows for simultaneous recording of Ca2+ signals and PER behavior. Even with the brain exposed to saline normal PER behavior was observed. Using the Gampi-Washi wick instead of a Kimwipe wick used in the original capillary method7 facilitates a highly reproducible and stable PER behavior and avoids the need to become skilled in making and choosing a good Kimwipe wick. The experimental tips stated above allowed us to successfully avoid disturbances by movement of the proboscis, leading to a very stable recording of Ca2+ signals with low noise. Occasionally, tissue removal was not sufficient to expose cells or suppress movement adequately, leading to poor results. However, once we were skilled, more than 80 % of preparations produced good results. This methodology is not only for Ca2+ imaging, but can also be adapted to any live imaging while observing PER behavior. For example, we can access any cell through the use of an electrode to directly record activity. Combined with two-photon excitation microscopy, we are able to perform time lapse imaging of synaptic structure, which can be related to behavioral changes. Therefore, this method of brain imaging with behavioral observation is not only valuable for the functional dissection of the neuronal network, but could be used as a powerful tool to correlate synaptic plasticity14 to the mechanisms behind memory.

<table border="1">
<tbody>
 <tr>
 <td>Name of Equipment/Reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Tetric EvoFlow</td>
 <td>Ivoclar vivadent</td>
 <td>M04115</td>
 <td>Light-curing glue</td>
 </tr>
 <tr>
 <td>BX51WI Microscope</td>
 <td>Olympus</td>
 <td>BX51WI</td>
 <td>With 40X (N.A. 0.8) water immersion objective lens</td>
 </tr>
 <tr>
 <td>Spinning disk confocal microscope system</td>
 <td>Improvision 			in PerkinElmer</td>
 <td>&nbsp;</td>
 <td>With 491 nm laser</td>
 </tr>
 <tr>
 <td>Stereomicroscope</td>
 <td>Nikon</td>
 <td>SMZ-800</td>
 <td>Attached to a swing arm. Stereo view is still 			available with video recording</td>
 </tr>
 <tr>
 <td>Gampi-Washi paper</td>
 <td>Haibara, Japan</td>
 <td>&nbsp;</td>
 <td>A special kind of traditional Japanese paper</td>
 </tr>
 <tr>
 <td>CCD camera</td>
 <td>Imaging Source</td>
 <td>DFK41AU02</td>
 <td>1/2" 1080 &times; 960 pixels SONY CCD</td>
 </tr>
 <tr>
 <td>Joystick manipulator</td>
 <td>Narishige</td>
 <td>MN-151</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Injector</td>
 <td>Narishige</td>
 <td>IM-5B</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster at JoVE.com

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.

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-recording-calcium-signals-from-identified-neurons

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Neuroscience

15.5K Views.  University of Massachusetts Medical School. ذبابة الفاكهة، ذبابة الفاكهة السوداء البطن، وتمتد خرطوم من أجل تغذية، واستجابة لحافز السكر من خرطوم الفيل، أو رسغ القدم. وقد جمعت أنا ملاحظات استجابة تمديد خرطوم الفيل (PER) مع الكالسيوم تقنية التصوير، مما يسمح لنا لرصد نشاط الخلايا العصبية في الدماغ، في ...

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

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

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 ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

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.

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 ...

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 ...

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 ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

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 ...

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To complicate matters even further, the odors detected by the insect nervous system usually are not single compounds but mixtures whose composition and concentration ratios vary. This leads to an almost infinite amount of different olfactory stimuli which have to be evaluated by the nervous system.
To understand which aspects of an odor stimulus determine its evaluation by the fly, it is therefore desirable to efficiently examine odor-guided behavior towards many odorants and odor mixtures. To directly correlate behavior to neuronal activity, behavior should be quantified in a comparable time frame and under identical stimulus conditions as in neurophysiological experiments. However, many currently used olfactory bioassays in Drosophila neuroethology are rather specialized either towards efficiency or towards resolution.
Flywalk, an automated odor delivery and tracking system, bridges the gap between efficiency and resolution. It allows the determination of exactly when an odor packet stimulated a freely walking fly, and to determine the animal&#180;s dynamic behavioral reaction.

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

The automated tracking system Flywalk is used for high-resolution quantification of odor-guided behavior in Drosophila melanogaster.

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To ...

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...