This work was supported by the German Research Council through the Collaborative Research Center SFB 889 &#34;Mechanisms of Sensory Processing&#34; and the Research Unit FOR 2705 &#34;Dissection of a Brain Circuit: Structure, Plasticity and Behavioral Function of the Drosophila Mushroom Body&#34;.

1. Transgenic fruit flies, Drosophila melanogaster
<ol>
	<li>Cross female virgin and male flies (raised at 25 &#176;C in 60% relative humidity on a 12 h light/dark cycle) carrying the desired Gal4 and UAS constructs25, respectively, to produce flies in which specific neurons of interest express a genetically encoded calcium indicator.</li>
	<li>Age the female progeny of the above cross until they are in the range of 3-6 days post-eclosion. Female flies are preferable because of their slightly larger size.</li>
</ol>
2. Preparation of the fruit fly for in vivo calcium imaging
<ol>
	<li>Select a single female fly and anesthetize on ice for no longer than 5 min.</li>
	<li>Using fine forceps, place the fly in the imaging chamber (demonstrated in Figure 1c). Ensure the thorax and legs are in contact with the electrical wires in the bottom of the chamber and that the head lays flat. Fix the position of the fly using clear adhesive tape. 
	NOTE: This protocol requires a custom-built chamber in which the flies are fixed, with the head capsule accessible for opening, and the flies able to receive both odor stimulations to the antennae and electric shocks to the thorax and legs (Figure 1).</li>
	<li>Using a surgical scalpel blade fixed to the scalpel handle (see Table of Materials), cut a window in the tape around the head of the fly, leaving the antennae covered and only the anterior-most portion of the thorax exposed.</li>
	<li>Surround the sides and back of the head with blue-light curing glue, carefully manipulated using an insect pin held by concave-convex jaws. Set the glue using a blue light-emitting LED lamp.</li>
	<li>Check that the glue is completely set and clear any residual unhardened glue from the dorsal surface of the fly head.</li>
	<li>Apply a drop of Ringer&#39;s solution24 (see Table of Materials) to cover the exposed cuticle of the head.</li>
	<li>Using a very fine-bladed stab knife, cut through the cuticle. For the most efficient removal of the cuticle, first cut across the posterior of the head using the ocelli as a starting point. Then, cut up each side, medial to the eyes, to form a flap of the cuticle that can be easily torn off using forceps.</li>
	<li>Remove any excess cuticle that may block the brain region of interest.</li>
	<li>Carefully clear the dorsal surface of any trachea using fine forceps, avoiding disruption of the brain tissue itself. Remove and refresh Ringer&#39;s solution24 (see Table of Materials) as required to clear the area of tissue debris.</li>
	<li>Place the hypodermic odor delivery needle in position, approximately 1 cm from the head of the fly. Ensure there is nothing that could obstruct odor delivery to the antennae.</li>
	<li>At the microscope, connect the imaging chamber to the odor-delivery system via the hypodermic odor delivery needle.</li>
	<li>To allow the fly to fully to recover from anesthesia and surgery, and to adapt to the air flow, implement a 10 min resting period at this stage.</li>
</ol>
3. In vivo calcium imaging
<ol>
	<li>Use a multiphoton microscope equipped with an infrared laser and a water immersion objective (see Table of Materials), installed on a vibration-isolated table. For the visualization of GFP-based calcium indicators, tune the laser to an excitation wavelength of 920 nm and install a GFP band-pass filter.</li>
	<li>Using the coarse Z adjustment knob, scan through the Z axis of the brain and locate the brain region of interest. Use the crop function to focus scanning on only this area to minimize scan time, and to rotate the scan view such that the anterior of the head is facing downwards.</li>
	<li>Adjust the frame size to 512 x 512 px, scan speed to &#62; 4 Hz, and scan region (in the Y dimension) so that the neurons of interest are covered.</li>
</ol>
4. Visualization of odor-evoked calcium transients through olfactory conditioning
<ol>
	<li>Use an odor delivery system19,26 that can deliver several odor stimuli in a temporally precise manner.
	<ol>
		<li>Use an additional computer to control the device, and to communicate with the imaging microscope software to coordinate odor stimulations with image capture during experiments.</li>
		<li>Initiate a pre-programmed macro package capable of linking the image acquisition software and odor delivery program (e.g., a VBA macro package installed in the microscope control software, see Table of Materials) that is responsive to an external input trigger provided by the initiation of an odor delivery protocol in a separate program).</li>
	</ol>
	</li>
	<li>Start the measurement by monitoring &#34;pre-training&#34;/na&#239;ve odor-evoked calcium transients at a resolution of 512 x 512 px and a frame rate of 4 Hz. Deliver a 2.5 s odor stimulus flanked by additional image acquisition for 6.25 s preceding odor onset (to establish an F0 baseline value) and 12.5 s after odor offset. Repeat this with a second odorant and then with a third odorant.</li>
	<li>Continue 3 min after this measurement with classical conditioning (&#34;training&#34;) the fly.
	<ol>
		<li>Select one of the odors presented in the &#34;pre-training&#34; phase to become the CS+ odor and another to become the CS- odor. Present the CS+ odor using the computer-controlled odor-deliver system for 60 s alongside twelve 90 V electric shocks.</li>
		<li>After a 60 s break, present the CS- odor alone for 60 s. Use as odorants 4-methylcyclohexanol and 3-octanol. Do not present the third odorant (e.g., 1-octen-3-ol) during this training as it is used as control only before (step 4.2.) and after (step 4.4) the training phase.</li>
	</ol>
	</li>
	<li>Measure the &#34;post-training&#34; odor-evoked calcium transients again by repeating the &#34;pre-training&#34; odor stimulation protocol (step 4.2.) 3 min after finishing the training phase (step 4.3). 
	NOTE: The timing of this step should reflect the time of interest after memory formation (e.g., carry out this step 3-4 min after the conditioning step to investigate short-term memory). Typically, flies can survive for several hours in this preparation.</li>
	<li>Save imaging files in an appropriate format (e.g., Tiff) for later image analysis.</li>
</ol>
5. Image analysis
<ol>
	<li>To analyze images, open Tiff (or similar) files in the image analysis software such as Fiji27.</li>
	<li>Align the stacks using a movement correction algorithm to ensure minimal movement in the X and Y directions. Discard any recordings that show strong movement in the Z axis.</li>
	<li>Select the tool of the software used to mark a region of interest (ROI) around the area that is to be examined. This should be as precise as possible to limit the influence of background fluorescence.</li>
	<li>Use the respective tool of the software to extract temporal fluorescence intensity data as data files from the selected ROI, such that a value is generated for each frame of the recording.</li>
	<li>Open the data using a data analysis program to calculate the &#916;F/F0 values for each odor trace. F0 = average fluorescence in the 2 s before odor onset. &#916;F = the difference between the raw fluorescence in a given frame and F0. From these values, calculate a &#916;F/F0 value for each frame which can be plotted against time to reflect relative fluorescence throughout the odor stimulus period.</li>
</ol>

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The authors have nothing to disclose.

The dissection of the neural circuitry underlying learning and memory is a prominent goal in the field of neuroscience. The genetic accessibility of Drosophila and the breadth and ease of behavioral testing makes this an ideal tool to investigate such phenomena. Here, a method is presented with which it is possible to visualize, within individual flies, the modulation that occurs at a subcellular level as a result of olfactory conditioning. By carrying out both pre-training and post-training visualization of odo...

Deciphering where and how the information is acquired in the brain through learning and subsequently stored as memory constitutes one of the most challenging tasks in neuroscience1. Neuroscientific research has led to the concept of a change in synaptic transmission as the neuronal substrate that underlies learning and memory formation2,3. It is hypothesized that, during learning, synaptic connections between neuronal ensembles that are active during the perception of a stimulus become modified such that their combined activity pattern can be retrieved during memory recall, thereby instructing future behavioral action4. These &#34;engram cells&#34; and their synapses are often distributed across brain regions and levels of processing, which makes it difficult to assign observed changes in synaptic transmission to the learning of a task or a stimulus. To localize and visualize those synaptic changes that are causally linked to a specific learning task one needs an appropriate model system that allows for precisely confining those synapses.
For such an endeavor, Drosophila melanogaster is particularly suitable because it combines relative brain simplicity, behavioral richness, and experimental accessibility. Among the well-established model organisms, Drosophila is situated between the nematode C. elegans and genetically tractable mammals like mice in terms of neuronal complexity. The stereotypic number of neurons (~300) and limited behavioral repertoire is observed in C. elegans. Mammals, on the other hand, have millions of neurons and staggering behavioral complexity. The brain of the fruit fly is, with its ~100,000, neurons significantly smaller than the brains of most vertebrates, and many of the neurons are individually identifiable5. Yet, Drosophila demonstrate a broad spectrum of complex behaviors, including an ability to exhibit robust associative olfactory learning and memory formation, first described over 40 years ago6. In the course of this classical conditioning procedure, groups of flies are subjected to an odor as the conditioned stimulus (CS+) while they receive a punishing electric shock as the unconditioned stimulus (US). A second odor (CS-) is then presented without any punishment. Thereby, the animals learn to avoid the odor associated with the punishment, which can be tested in a subsequent choice situation between the two odors, CS+ and CS-. Work on dissecting the neuronal substrate underlying this behavior in Drosophila has identified the mushroom bodies (MB) as the primary site of the &#34;engram&#34;7,8,9,10 and, therefore, the circuitry of this brain region was and is the subject of intense research in order to uncover the logic by which a memory engram is acquired and stored (recently reviewed in11,12).
The Drosophila MB consists of ~2,000 intrinsic neurons (Kenyon cells) per hemisphere, organized in parallel axonal projections13. Axons of olfactory projection neurons are extended to the lateral protocerebra and to the MB calyces, the main dendritic input site of the MB and receive olfactory input from antennal lobes. The long, parallel axons bundle of Kenyon cells constitute the peduncle and the lobes. Most Kenyon cells bifurcate forming horizontal &#946;/&#946;&#39;-lobes by extending one collateral towards the midline of the brain, and the vertical &#945;/&#945;&#39;-lobes by extending second collateral projecting in the dorsal-anterior direction. The other group of Kenyon cells forms the horizontal &#947;-lobes13 of the MB where the learning process and subsequent short-term memory formation could be localized10. The MB lobes receive afferent input and provide efferent output, both of which are typically restricted to distinct compartmental sub-regions along the Kenyon cell axons14,15,16. In particular, afferent dopaminergic MB input neurons have been shown to mediate value-based, e.g., punitive, reinforcing effects in associative olfactory learning15,17. Stereotypic and individually identifiable efferent MB output neurons from the mushroom body lobes integrate information across large numbers of Kenyon cells, target diverse brain areas and bear behavior-instructive appetitive or aversive information15. This neuronal architecture has led to a concept of the organization of the associative engram. Odors are relatively precisely encoded by sparsely activated ensembles of Kenyon cells. The coincident activity of these Kenyon cell ensembles and release of dopamine - evoked by punishing stimuli - modulates transmission from Kenyon cell presynapses onto MB output neurons such that the animals will subsequently avoid this particular smell10,12. We use this rather precisely defined and localized engram as a paradigmatic case to illustrate how these learning-dependent changes in synaptic activity can be determined and monitored.
The value of Drosophila as a model system relies strongly on the unmatched genetic toolbox that allows one to express transgenes for identifying, monitoring, and controlling single neurons within complex circuits18. The advent of techniques for neuronal activity monitoring - such as calcium imaging, discussed here - have allowed for the determination of neuronal activity patterns in response to a specific stimulus. By combining specific Gal4-driven expression of genetically encoded calcium indicators (GECIs) with olfactory stimulation, one can visualize the odor-evoked calcium dynamics of neurons of interest19. In this protocol, it is shown that by further coupling this technique with a classical conditioning paradigm, it is possible to examine these olfactory responses in the context of learning. Learning-induced plasticity can be further dissected using GECIs that are not only localized to a single specific neuron, but also to specific subcompartments of a neuron. Pech et al.20 established a selection of tools that allow exactly this. By targeting GCaMP321 to either the pre- or postsynapse - via linkage to the vertebrate Synaptophysin or dHomer, respectively20- the differential modulation of these sites can be distinguished. This localization confers, in this context, an advantage over most GECIs that are ubiquitously present throughout the cytosol - e.g., GCaMP22, GCaMP321, or GCaMP623 - because it means that pre- and postsynaptic transients can be distinguished from the overall integrated calcium influx that occurs as a result of neuron activation. This can provide clues about the location and types of plasticity that occur as a result of or that cause learning and memory formation. As an example, the protocol provided here shows the value of this tool in deciphering the modulation of MB output neurons during olfactory associative learning by targeting the expression of the calcium sensor to only the postsynapse. By monitoring, within an individual fly, odor-evoked activity before and after olfactory conditioning a direct comparison can be drawn between a na&#239;ve odor response and a learned odor response. Whilst fixed in the same imaging chamber, flies are exposed to a selection of odors. Then, they receive an aversive associative conditioning protocol in which one of these odors is paired with electric shock (becoming the CS+) and another odor is presented without reinforcement (becoming the CS-). Finally, the flies are again exposed to the same odors as in the first step. Calcium dynamics are observed using two-photon microscopy.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1-Octen-3-ol</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>O5284</td>
			<td>Chemical used as odorant</td>
		</tr>
		<tr>
			<td>3-Octanol</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>218405</td>
			<td>Chemical used as odorant</td>
		</tr>
		<tr>
			<td>4-Methylcyclohexanol</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>153095</td>
			<td>Chemical used as odorant</td>
		</tr>
		<tr>
			<td>Bandpass filter for EGFP (525/50 nm)</td>
			<td>Carl Zeiss Microscopy GmbH, Jena, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Clear adhesive tape</td>
			<td>Tesa SE, Norderstedt, Germany</td>
			<td></td>
			<td>Standard claer adhesive tape</td>
		</tr>
		<tr>
			<td>Concave-convex jaws</td>
			<td>Fine Science Tools, North Vancouver, Canada</td>
			<td>10053-09</td>
			<td>Blade Holders with concave-convex jaws</td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Fine Science Tools, North Vancouver, Canada</td>
			<td>11412-11</td>
			<td>Forceps with tip 0.1 x 0.06mm</td>
		</tr>
		<tr>
			<td>Hypodermic needle</td>
			<td>Sterican - B. Braun, Melsungenk, Germany</td>
			<td>4665120</td>
			<td>1.20x40mm</td>
		</tr>
		<tr>
			<td>Insect Minutien pins</td>
			<td>Fine Science Tools, North Vancouver, Canada</td>
			<td>26002-10</td>
			<td>Diameter 0.1mm, tip 0.0125mm</td>
		</tr>
		<tr>
			<td>Kentoflow</td>
			<td>Kent Express Dental Supplies, Gillingham, UK</td>
			<td>953683</td>
			<td>Blue light-curing glue</td>
		</tr>
		<tr>
			<td>Microscope slide</td>
			<td>Carl Roth GmbH &#38; Co. KG, Karlsruhe, Germany</td>
			<td>0656.1</td>
			<td>Standard objective slide 76 x 26 mm</td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>M8410</td>
			<td>Used as diluent for odorants</td>
		</tr>
		<tr>
			<td>Mode-locked Ti-Sapphire laser Chameleon Vision 2</td>
			<td>Coherent Inc., Santa Clara, CA, USA</td>
			<td></td>
			<td>Tunable infrared femtosecond laser</td>
		</tr>
		<tr>
			<td>Multiphoton Microscope LSM 7MP equipped with BiG detectors</td>
			<td>Carl Zeiss Microscopy GmbH, Jena, Germany</td>
			<td></td>
			<td>Multiphoton microscope, multiple companies provide similar devices.</td>
		</tr>
		<tr>
			<td>Plan-Apochromat 20x (NA = 1.0) water immersion objective</td>
			<td>Carl Zeiss Microscopy GmbH, Jena, Germany</td>
			<td>421452-9900-000</td>
			<td>Objective W &#34;Plan-Apochromat&#34; 20x/1.0 DIC M27 70mm</td>
		</tr>
		<tr>
			<td>Ringer&#39;s solution</td>
			<td>n.a.</td>
			<td>n.a.</td>
			<td>5mM KCl, 130mM NaCl, 2mM MgCl2, 2mM CaCl2, 5mM Hepes-NaOH, 36mM sucrose, pH = 7.4</td>
		</tr>
		<tr>
			<td>Stab knife</td>
			<td>Sharpoint, Surgical Specialties Corporation, Reading, PA, USA</td>
			<td>72-1551</td>
			<td>5.0mm Straight restricted blade depth</td>
		</tr>
		<tr>
			<td>Surgical scalpel blade</td>
			<td>Swann-Morton, Sheffield, UK</td>
			<td>0303</td>
			<td>Product No. 11</td>
		</tr>
		<tr>
			<td>Surgical scalpel handle</td>
			<td>Swann-Morton, Sheffield, UK</td>
			<td>0907</td>
			<td>Product No. 7S/S</td>
		</tr>
		<tr>
			<td>Visual Basics of Applicatons (VBA) software to receive a trigger 
			from the odor-delivery device and the electric shock 
			application device (power supply) to interact with the 
			ZEN software from Zeiss that controls the microscope.</td>
			<td>Custom-written and available upon request</td>
			<td>n.a.</td>
			<td>n.a.</td>
		</tr>
	</tbody>
</table>

in vivo optical calcium imaging of learning-induced synaptic plasticity in drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

An example of images acquired with the above protocol can be seen in Figure 2. dHomer-GCaMP3 is expressed in an MB output neuron whose dendrites innervate the compartment 1 of the MB &#947;-lobe (the neuron is termed MVP228,29) and is genetically targeted using the split-Gal4 line MB112C16. Also, demonstrated is the difference in the subcellular localization of a cytos...

Watch this Scientific Journal Video about In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster at JoVE.com

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. ...

in-vivo-optical-calcium-imaging-learning-induced-synaptic-plasticity

Research

JoVE Journal

Neuroscience

8.8K Views.  University of Göttingen. Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

Video: In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new method of Ca2+-imaging using the bioluminescent reporter GFP-aequorin (GA) has been developed. This new technique relies on the fusion of the GFP and aequorin genes, producing a molecule capable of binding calcium and&#160;&#8212; with the addition of its cofactor coelenterazine &#8212; emitting bright light that can be monitored through a photon collector. Transgenic lines carrying the GFP-aequorin gene have been generated for both mice and Drosophila. In Drosophila, the GFP-aequorin gene has been placed under the control of the GAL4/UAS binary expression system allowing for targeted expression and imaging within the brain. This method has subsequently been shown to be capable of detecting both inward Ca2+-transients and Ca2+-released from inner stores. Most importantly it allows for a greater duration in continuous recording, imaging at greater depths within the brain, and recording at high temporal resolutions (up to 8.3 msec). Here we present the basic method for using bioluminescent imaging to record and analyze Ca2+-activity within the mushroom bodies, a structure central to learning and memory in the fly brain.

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Here we present a novel Ca2+-imaging approach using a bioluminescent reporter. This approach uses a fused construct GFP-aequorin which binds to Ca2+ and emits light, eliminating the need for light excitation. Significantly this method permits long continuous imaging, access to deep brain structures and high temporal resolution.

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new ...

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at the synaptic level. The Active Zone (AZ) is the region at the presynaptic membrane that mediates neurotransmitter release and is composed of a dense collection of scaffold proteins. AZs of Drosophila melanogaster (Drosophila) photoreceptors undergo molecular remodeling after prolonged exposure to natural ambient light. Thus the level of neuronal activity can rearrange the molecular composition of the AZ and contribute to the regulation of the functional output.
Starting from the light exposure set-up preparation to the immunohistochemistry, this protocol details how to quantify the number, the spatial distribution, and the delocalization level of synaptic molecules at AZs in Drosophila photoreceptors. Using image analysis software, clusters of the GFP-fused AZ component Bruchpilot were identified for each R8 photoreceptor (R8) axon terminal. Detected Bruchpilot spots were automatically assigned to individual R8 axons. To calculate the distribution of spot frequency along the axon, we implemented a customized software plugin. Each axon&#39;s start-point and end-point were manually defined and the position of each Bruchpilot spot was projected onto the connecting line between start and end-point. Besides the number of Bruchpilot clusters, we also quantified the delocalization level of Bruchpilot-GFP within the clusters. These measurements reflect in detail the spatially resolved synaptic dynamics in a single neuron under different environmental conditions to stimuli.

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

Here we show how to quantify the number and spatial distribution of synaptic active zones in Drosophila melanogaster&#160;photoreceptors, highlighted with a genetically encoded molecular marker, and their modulation after prolonged exposure to light.

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at ...

Slice Patch Clamp Technique for Analyzing Learning-Induced Plasticity

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze motor-learning induced plasticity, we trained rats using an accelerated rotor rod task. Rats performed the task 10 times at 30-s intervals for 1 or 2 days. Performance was significantly improved on the training days compared to the first trial. We then prepared acute brain slices of the primary motor cortex (M1) in untrained and trained rats. Current-clamp analysis showed dynamic changes in resting membrane potential, spike threshold, afterhyperpolarization, and membrane resistance in layer II/III pyramidal neurons. Current injection induced many more spikes in 2-day trained rats than in untrained controls.
To analyze contextual-learning induced plasticity, we trained rats using an inhibitory avoidance (IA) task. After experiencing foot-shock in the dark side of a box, the rats learned to avoid it, staying in the lighted side. We prepared acute hippocampal slices from untrained, IA-trained, unpaired, and walk-through rats. Voltage-clamp analysis was used to sequentially record miniature excitatory and inhibitory postsynaptic currents (mEPSCs and mIPSCs) from the same CA1 neuron. We found different mean mEPSC and mIPSC amplitudes in each CA1 neuron, suggesting that each neuron had different postsynaptic strengths at its excitatory and inhibitory synapses. Moreover, compared with untrained controls, IA-trained rats had higher mEPSC and mIPSC amplitudes, with broad diversity. These results suggested that contextual learning creates postsynaptic diversity in both excitatory and inhibitory synapses at each CA1 neuron.
AMPA or GABAA receptors seemed to mediate the postsynaptic currents, since bath treatment with CNQX or bicuculline blocked the mEPSC or mIPSC events, respectively. This technique can be used to study different types of learning in other regions, such as the sensory cortex and amygdala.

The slice patch clamp technique is an effective method for analyzing learning-induced changes in the intrinsic properties and plasticity of excitatory or inhibitory synapses.

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze ...

Pentylenetetrazole-Induced Kindling Mouse Model

Pentylenetetrazole (PTZ) is a GABA-A receptor antagonist. An intraperitoneal injection of PTZ into an animal induces an acute, severe seizure at a high dose, whereas sequential injections of a subconvulsive dose have been used for the development of chemical kindling, an epilepsy model. A single low-dose injection of PTZ induces a mild seizure without convulsion. However, repetitive low-dose injections of PTZ decrease the threshold to evoke a convulsive seizure. Finally, continuous low-dose administration of PTZ induces a severe tonic-clonic seizure. This method is simple and widely applicable to investigate the pathophysiology of epilepsy, which is defined as a chronic disease that involves repetitive seizures. This chemical kindling protocol causes repetitive seizures in animals. With this method, vulnerability to PTZ-mediated seizures or the degree of aggravation of epileptic seizures was estimated. These advantages have led to the use of this method for screening anti-epileptic drugs and epilepsy-related genes. In addition, this method has been used to investigate neuronal damage after epileptic seizures because the histological changes observed in the brains of epileptic patients also appear in the brains of chemical-kindled animals. Thus, this protocol is useful for conveniently producing animal models of epilepsy.

This protocol describes a method of chemical kindling with pentylenetetrazole and provides a mouse model of epilepsy. This protocol can also be used to investigate vulnerability to seizure induction and pathogenesis after epileptic seizures in mice.

Pentylenetetrazole (PTZ) is a GABA-A receptor antagonist. An intraperitoneal injection of PTZ into an animal induces an acute, severe seizure at a high ...

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model animal for endocrine research, Drosophila melanogaster, which has sophisticated genetic tools and genome information, is being increasingly used. This article describes a method for the calcium imaging of Drosophila brain explants. This method enables the detection of the direct signaling of a hormone to the brain. It is well known that many peptide hormones act through G-protein-coupled receptors (GPCRs), whose activation causes an increase in the intracellular Ca2+concentration. Neural activation also elevates intracellular Ca2+ levels, from both Ca2+ influx and the release of Ca2+ stored in the endoplasmic reticulum (ER). A calcium sensor, GCaMP, can monitor these Ca2+ changes. In this method, GCaMP is expressed in the neurons of interest, and the GCaMP-expressing larval brain is dissected and cultured ex vivo. The test peptide is then applied to the brain explant, and the fluorescent changes in GCaMP are detected using a spinning disc confocal microscope equipped with a CCD camera. Using this method, any water-soluble molecule can be tested, and various cellular events associated with neural activation can be imaged using the appropriate fluorescent indicators. Moreover, by modifying the imaging chamber, this method can be used to image other Drosophila organs or the organs of other animals.

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

This paper describes a protocol for ex vivo calcium imaging of the Drosophila brain. In this method, natural or synthetic compounds can be applied to the buffer to test their ability to activate particular neurons in the brain.

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

3D Modeling of Dendritic Spines with Synaptic Plasticity

Computational modeling of diffusion and reaction of chemical species in a three-dimensional (3D) geometry is a fundamental method to understand the mechanisms of synaptic plasticity in dendritic spines. In this protocol, the detailed 3D structure of the dendrites and dendritic spines is modeled with meshes on the software Blender with CellBlender. The synaptic and extrasynaptic regions are defined on the mesh. Next, the synaptic receptor and synaptic anchor molecules are defined with their diffusion constants. Finally, the chemical reactions between synaptic receptors and synaptic anchors are included and the computational model is solved numerically with the software MCell. This method describes the spatiotemporal path of every single molecule in a 3D geometrical structure. Thus, it is very useful to study the trafficking of synaptic receptors in and out of the dendritic spines during the occurrence of synaptic plasticity. A limitation of this method is that the high number of molecules slows the speed of the simulations. Modeling of dendritic spines with this method allows the study of homosynaptic potentiation and depression within single spines and heterosynaptic plasticity between neighbor dendritic spines.

The protocol develops a three-dimensional (3D) model of a dendritic segment with dendritic spines for modeling synaptic plasticity. The constructed mesh can be used for computational modeling of AMPA receptor trafficking in the long-term synaptic plasticity using the software program Blender with CellBlender and MCell.

Computational modeling of diffusion and reaction of chemical species in a three-dimensional (3D) geometry is a fundamental method to understand the ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.