SG is supported by funds from the State University of New York at Buffalo and from John R. Oishei Foundation.
MK was supported by a Student Research Award by CURCA at UB.

1. Preparation of Reagents
<ol>
	<li>Prepare the dissection buffer preparation by adding the following compounds: 128 mM NaCl, 1mM EGTA, 4 mM MgCl2, 2mM KCl, 5mM HEPES, and 36 mM Sucrose in 1000 mL, PH to 7.2 and filter sterilize.</li>
</ol>
2. Preparation for the Dissection
<ol>
	<li>Siligard cube: The gel is cut to approximately one inch wide, 1 inch long and about 1cm high. The gel cube is placed on a glass slide during dissection. Cubes may be used more than once.</li>
	<li>Pins: Pins used for dissection need to be extra short. The sharp bottom portion of minutien pins works best in dissection.</li>
	<li>Need one sharp microdissection spring scissors.</li>
	<li>Need two fine tipped forceps.</li>
</ol>
3. Collection of Larvae
<ol>
	<li>Wandering 3rd instar larvae that are expressing a GFP tagged vesicle protein are collected onto a petri dish.</li>
	<li>Larvae are washed in deionized water to get rid of all the food.</li>
</ol>
4. Live dissection of larvae
<ol>
	<li>A larva is transferred to the dissection gel cube on a glass slide. Larva is immersed in dissection buffer.</li>
	<li>Larva is pined at the anterior and posterior using two fine pins with the dorsal side up.</li>
	<li>Using microdissection scissors, a small cut is made at the midline near the posterior end. From this incision a cut is made through to the anterior. Using two pins, pin the cut cuticle on the lateral edges.</li>
	<li>Add a drop of dissecting buffer. Carefully remove the gut, intestines and fat bodies with forceps. Most of these tissues normally ooze out during the dorsal cutting.</li>
	<li>Replace the buffer with fresh dissecting buffer. The larva should never be dry and should constantly be in dissecting buffer.</li>
	<li>Incubations using in vivo markers MitoTracker or LysoTracker are done at this time, by diluting the in vivo marker in dissection buffer. After the incubation of the in vivo marker wash the dissected larva several times (5 quick washes) using fresh dissecting buffer.</li>
	<li>After replacing the last wash push the pins into the syligard and immediately coverslip. The larvae is now ready for observation.</li>
</ol>
5. Live imaging of dissected larvae
<ol>
	<li>Place the coverslip and gel onto the microscope stage and image using an inverted microscope.</li>
	<li>We use a 100x lens to image tagged vesicles within larval segmental nerves. GFP expression in the cell bodies of the ventral ganglion is first evaluated before segmental nerves are imaged. If robust GFP staining is observed in the cell bodies in the ventral ganglion, the nerves are then imaged and used for evaluation.</li>
	<li>Using a dual view system with filters for CFP/YFP or RFP/YFP and split view software we can simultaneously image two tagged protein in vivo.</li>
</ol> 
6. Representative Results
<img src="/files/ftp_upload/2151/2151fig1.jpg" alt="Figure 1" /> Figure 1: A representative image showing many axonal vesicles on many axonal tracks within a larval segmental nerve. Note the blobs of GFP, representing axonal accumulations.
<img src="/files/ftp_upload/2151/2151fig2.jpg" alt="Figure 2" /> Figure 2: A representative image showing a single axonal track with many axonal vesicles within a larval segmental nerve.

<ol>
	<li>Brand, A. H., Perrimon, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+gene+expression+as+a+means+of+altering+cell+fates+and+generating+dominant+phenotypes.">Targeted gene expression as a means of altering cell fates and generating dominant phenotypes.</a> Development. 118, 401-415 (1993).</li><li>Hurd, D. D., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinesin+mutations+cause+motor+neuron+disease+phenotypes+by+disrupting+fast+axonal+transport+in+Drosophila.">Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila.</a> Genetics. 144, 1075-1085 (1996).</li><li>Gunawardena, S., Her, L. S., Brusch, R. G., Laymon, R. A., Niesman, I. R., Gordesky-Gold, B., Sintasath, L., Bonini, N. M., Goldstein, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+axonal+transport+by+loss+of+huntingtin+or+expression+of+pathogenic+polyQ+proteins+in+Drosophila.">Disruption of axonal transport by loss of huntingtin or expression of pathogenic polyQ proteins in Drosophila.</a> Neuron. 40, 25-40 (2003).</li></ol>

No conflicts of interest declared.

In vivo imaging of synaptic vesicle transport within Drosophila larval segmental nerves is a powerful tool for studying the mechanisms in axonal transport. We have previously used this protocol to evaluate the transport dynamics within larval nerves expressing expanded polyQ repeats to elucidate how expansion of repeats affect the dynamics of vesicle transport 3. Using this protocol the velocity of vesicle movement can be determined by converting the video streams to a kymograph. Using geneti...

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		<div>
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		<thead>
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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
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		</thead>
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<tr>
<td>1 Pair Micro Dissection Scissors</td>
<td>&nbsp;</td>
<td>Fine Scientific </td>
<td>&nbsp;</td>
<td>Spring scissors &#x2013; 6mm blade. 0.125 mm tip diameter</td>
</tr>
<tr>
<td>2 Pair Forceps</td>
<td>&nbsp;</td>
<td>Fischer Scientific</td>
<td>&nbsp;</td>
<td>Fisherbrand Dissecting Micro-Adson serrated tip forceps</td>
</tr>
<tr>
<td>Box of Dissection Pins</td>
<td>&nbsp;</td>
<td>Fine Scientific </td>
<td>&nbsp;</td>
<td>Minutien Pins 0.1mm diameter</td>
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in vivo visualization of synaptic vesicles within drosophila larval segmental axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

Watch this Scientific Journal Video about In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons at JoVE.com

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

in-vivo-visualization-synaptic-vesicles-within-drosophila-larval

Research

JoVE Journal

Neuroscience

9.4K Views.  SUNY-University at Buffalo. Questo protocollo descrive la dissezione dal vivo di Drosophila allo scopo di immagini del movimento di GFP vescicole tag assonale sulle piste microtubuli.

Video: In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Laser Nanosurgery of Cerebellar Axons In Vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to understand the mechanism that promotes neuronal regeneration, an in-depth analysis of the neuronal types that can remodel after injury is needed. Several studies showed that damaged climbing fibers are capable of regrowing also in adult animals1,2. The investigation of the time-lapse dynamics of degeneration and regeneration of these axons within their complex environment can be performed by time-lapse two-photon fluorescence (TPF) imaging in vivo3,4. This technique is here combined with laser surgery, which proved to be a highly selective tool to disrupt fluorescent structures in the intact mouse cortex5-9.
This protocol describes how to perform TPF time-lapse imaging and laser nanosurgery of single axonal branches in the cerebellum in vivo. Olivocerebellar neurons are labeled by anterograde tracing with a dextran-conjugated dye and then monitored by TPF imaging through a cranial window. The terminal portion of their axons are then dissected by irradiation with a Ti:Sapphire laser at high power. The degeneration and potential regrowth of the damaged neuron are monitored by TPF in vivo imaging during the days following the injury.

Laser Nanosurgery of Cerebellar Axons In Vivo

Two-photon imaging, coupled to laser nanodissection, are useful tools to study degenerative and regenerative processes in the central nervous system with subcellular resolution. This protocol shows how to label, image, and dissect single climbing fibers in the cerebellar cortex in vivo.

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Culturing In Vivo-like Murine Astrocytes Using the Fast, Simple, and Inexpensive AWESAM Protocol

The AWESAM (a low-cost easy stellate astrocyte method) protocol entails a fast, simple, and inexpensive way to generate large quantities of in vivo-like mouse and rat astrocyte monocultures: Brain cells can be isolated from different brain regions, and after a week of cell culture, non-astrocytic cells are shaken off by placing the culture dishes on a shaker for 6 h in the incubator. The remaining astrocytes are then passaged into new plates with an astrocyte-specific medium (termed NB+H). NB+H contains low concentrations of heparin-binding EGF-like growth factor (HBEGF), which is used in place of serum in medium. After growing in NB+H, AWESAM astrocytes have a stellate morphology and feature fine processes. Moreover, these astrocytes have more in vivo-like gene expression than astrocytes generated by previously published methods. Ca2+ imaging, vesicle dynamics, and other events close to the membrane can thus be studied in the fine astrocytic processes in vitro, e.g., using live cell confocal or TIRF microscopy. Notably, AWESAM astrocytes also exhibit spontaneous Ca2+ signaling similar to astrocytes in vivo.

Culturing In Vivo-like Murine Astrocytes Using the Fast, Simple, and Inexpensive AWESAM Protocol

The AWESAM protocol described here is optimal for culturing murine astrocytes in isolation from other brain cells in a fast, simple, and inexpensive manner. AWESAM astrocytes exhibit spontaneous Ca2+ signaling, morphology, and gene expression profiles similar to astrocytes in vivo.

The AWESAM (a low-cost easy stellate astrocyte method) protocol entails a fast, simple, and inexpensive way to generate large quantities of in vivo-like ...

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal macropatch recordings of synaptic currents from visualized boutons at the Drosophila larval NMJ. This technique requires customized fabrication of recording micropipettes, as well as a compound microscope equipped with a high magnification, long-distance water immersion objective, differential interference contrast (DIC) optics, and a fluorescent attachment. The recording electrode is positioned on the top of a selected synaptic bouton visualized with DIC optics, epi-fluorescence, or both. The advantage of this technique is that it allows monitoring the synaptic activity of a limited number of sites of release. The recording electrode has&#160;a diameter of several microns, and the release sites positioned outside of the electrode rim do&#160;not significantly affect the recorded currents. The recorded synaptic currents have fast kinetics and can be readily resolved. These advantages are especially important for the studies of mutant fly lines with enhanced spontaneous or asynchronous synaptic activity.

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Synaptic currents can be recorded focally from visualized synaptic boutons at the Drosophila third instar larvae neuromuscular junction. This technique enables monitoring the activity of a single synaptic bouton.

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

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.

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.