This work was supported by DoD Grant PT074620 (SLJ) and NIH Grant R01NS245014 (SLJ).

1. Reagent preparation
<ol>
 <li>Thaw frozen Matrigel on wet ice, leaving enough time before the experiment (do not speed up the process by active warming). Matrigel is used to establish a 3D environment closely resembling an extracellular matrix conducive to the migration of cells. It provides cells with both a substrate and mechanical encasing. Matrigel is stored frozen at -20&deg;C. Prior to use, it should be gradually brought up to the temperature of 0-4&deg;C, i.e. thawed on wet ice or in a refrigerator. Matrigel irreversibly sets to form a gel in the temperature of incubation (37&deg;C). Therefore it is not recommended to speed up thawing by immersing the container in warm water.</li>
 <li>Protein fractions are prepared by isoelectric focusing (IEF) of the total protein extract from the cerebral cortex of postnatal day 0 (P0) ferret. After separation, the fractions are dialyzed overnight against 20mM Tris-HCl pH7.2 to clean the extracts from the focusing buffer constituents, which might be toxic to the tissue.</li>
 <li>Protein delivery system. Proteins extracted from the cerebral cortex are adsorbed on fluorescent latex beads which then slowly release the proteins to the environment.We found that the fluorescence feature helps in visual handling of microspheres and allows easy detection of even minimal spillovers from the site of bead deposit, when observed under fluorescent microscope.
 <ol class="lalpha">
 <li>Add 10&mu;l of fluorescent beads to 100-200&mu;l of protein solution</li>
 <li>Incubate for 30min in dark at RT on a shaker</li>
 <li>Spin the beads down in a bench centrifuge for 1hr at max speed</li>
 <li>Remove supernatant</li>
 <li>Re-suspend in a vortex using 10&mu;l of fresh medium</li>
 <li>Add 30&mu;l of Matrigel to the suspension, mix well, and place on ice.</li>
 </ol>
 </li>
<li>Buffers and media

<ol>
 <li>Prepare and filtration-sterilize artificial cerebrospinal fluid (aCSF; NaCl 124mM, NaHCO3 26mM, NaH2PO4 1.2mM, KCl 3.2mM, MgSO4 1.2mM, CaCl2 2.4mM, Glucose 10mM). Slush-freeze the aCSF beforehand, so it's ready at the time of tissue dissection. To make it, place a plastic flask filled with sterile aCSF in a -80&deg;C freezer. Monitor freezing every so often, so when a layer of ice builds up on the inside of wall, break it and mix with the fluid. Repeat until the flask contains mostly an icy slush. Buffer prepared this way not only supports tissue survival, but will also hold low temperature longer, and is mechanically less harmful to the tissue than chunky ice cubes.</li>
 <li>Prepare Neurobasal medium with N-2 and B-27 supplements added according to manufacturer's suggestions, as well as 1mg/100ml gentamycin, 2 mM L-glutamine, and 0.6 % glucose. Sterilize by filtration through a 0.22&mu;m membrane. Keep warm at 37&deg;C until use.</li>
</ol>
</li>
<li>Culture plates
 <ol class="lalpha">
 <li>Culture grade plastic six-well plates are prepared by dotting each well in its center with a small amount (about 2&mu;l) of cold Matrigel using 20&mu;l pipette tip, and placing an autoclaved round 18mm glass coverslip on each drop. Neither the plates nor the coverslips need to be otherwise culture treated. </li>
 </ol>
</li>
</ol>
2. Tissue preparation
<ol>
 <li>Obtaining the brain tissue
 <ol class="lalpha">
 <li>After deeply anesthetizing the animal with Euthasol, remove the brain and place it in ice-cold aCSF</li>
 <li>Separate the hemispheres with a scalpel and slice coronally at 500&mu;m using a device of choice, collecting slices into a small dish containing ice-cold aCSF</li>

 </ol>
 
 </li>
 
 <li>Making explants (use dissecting microscope)
 <ol class="a">
 <li>Identify slices containing the ganglionic eminences (GE) and trim regions of cortex that may curl over GE and interfere with further steps</li>
 <li>Press a Harris Uni-Core bore in the area close to ventricular surface of GE, retract a core containing predominantly a ventricular zone, and expel it with a plunger into the ice-cold medium (Fig.1). Three to four explants of 0.5 mm diameter can be made from the GE in single P0 ferret brain slice. Please note that in both rodents and ferrets, the medial and lateral GEs are fused at this age. If a core stays in the slice or is stuck to the plastic bottom of the dish, mobilize it by squirting aCSF with a pipette tip and transfer with a pipette into the dish containing cold Neurobasal medium. Keep the dish on wet ice.</li>
 <li>Transfer a batch of explants from the medium into a large drop of ice-cold Matrigel (about 0.25-0.5ml) placed in a separate dish, taking care to have the explants fully immersed and avoiding bubbles. Keep the dish on wet ice. The size of a batch depends on the speed of the workflow downstream.</li>
 
 </ol> 
 </li>
</ol>
3. Culture setup (use dissecting microscope)
<ol>
 <li>Always keep Matrigel and Matrigel-containing mixes on ice to avoid curing.</li>
 <li>Remix the bead suspension and place 1-2&mu;l or less of the bead suspension in ice-cold Matrigel in the center of a glass coverslip previously placed in a culture plate well using a 20&mu;l pipette tip. Let it set a little, avoid drying.</li>
 <li>Using a 20&mu;l pipette tip pick up 2 explants from the Matrigel drop and place them in the desired vicinity of the bead deposit (at least 1mm away from the edge of the deposit) symmetrically on the opposite sides. Let it set for no longer than 5min at room temperature. The right timing of steps b and c is critical. If the time is too short it will lead to incomplete gelling and components of the culture setup will move when they are covered with the final layer of Matrigel; if the time is too long it may result in over-drying of the gel and creation of physical barriers impeding the free movement of cells.</li>
<li>Cover the beads and the explant with a flat layer of about 30&mu;l of Matrigel, extending it beyond the explants (Fig.2). Let it set in the incubator for about 15 minutes.</li>
<li>Add 2ml of Neurobasal medium with supplements.</li>
<li>Incubate at 37&deg;C in 5% CO2/95% O2 monitoring migratory activity daily.</li>
</ol>
4. Live imaging
<ol>
 <li>After incubation for 3-4 days, review the culture plates and choose coverslips of interest. The explants should generate a radially uniform cloud of cells. Replace chosen coverslips into a fresh culture plate that will be used in live imaging.</li>
<li>Let the coverslips sit in the new plate over night. Before placing the plate on the microscope for imaging, in a cell culture hood, use a sterile spatula to move the coverslip in order to release any microbubbles that collect between the coverslip and the cell culture plate. Interference of bubbles will ruin the images. Move the coverslips to the center of the well.</li>
<li>To prevent evaporation, fill the space between wells with water.</li>
<li>Carefully transfer the plate from the cell culture hood to the microscope as to not move the coverslips from the center of the well.</li>
<li>Define an imaging scheme using imaging software. Due to the 3D nature of the culture setup the cells also move vertically, therefore it is important to collect Z-planes encompassing full depth of the migration cloud. The XY extent of the imaged area should also be set large enough to cover areas predicted to be populated by migrating cells during the course of recording, and to contain the bead deposit in order to record an angular orientation of the explant and migrating cells relative to the source of the protein. Also consider that the more imaged Z planes, or the more area covered, the more time it takes to record a cycle at a particular time point, forcing longer time intervals between samples and a loss of temporal resolution of cell tracking. In our case, using a 10x objective, the optimal scheme was a 3x3 tile recording for each of 10 Z-planes spaced every 20&mu;m, at intervals of 30min. We found 30 min intervals the longest that allows unequivocal identification of cells at consecutive time points, which is critical for reliable tracking of their movements.</li>
<li>Initiate time lapse imaging.</li>
<li>After the desired time, remove the plate and fix cultures in 4% phosphate-buffered paraformaldehyde. It can be used for further morphological and immunocytochemical analysis.</li>
</ol>
5. Analysis of movement
<ol>
 <li>Convert the recorded time series of images into a format most convenient for further processing, TIFF being the most portable one.</li> 
 <li>Track the movement of individual cells over time by recording XY coordinates. This can be done using freely available software like ImageJ (NIH) with a Manual Tracking plug-in, or any other commercial application such as ImagePro, Volicity or AutoQuant.</li>
 <li>Plot the tracks using Excel or in-house developed software.</li>
 <li>For each tracked cell calculate kinetic parameters such as average speed, curvature of path, directional variability over time, branching, deviation toward the source of proteins, etc. Calculations can be done in MS Excel or any equivalent spreadsheet. Our experience indicates however that it is far more productive to develop in-house scripts (using R, Matlab, Python, Perl or other, mostly Open Source languages), as they allow for more flexible customization and inclusion of more sophisticated analytical methods than those available in a basic installation of Excel.</li>

</ol>

6. Representative Results:

<img src="/files/ftp_upload/2583/2583fig1.jpg" alt="Figure 1"/> 
Figure 1. Diagram of the anatomical origin of explants. This is a drawing of a coronal slice of a hemisected ferret brain. The grey circles indicate the region from which the explants were taken. 
<img src="/files/ftp_upload/2583/2583fig2.jpg" alt="Figure 2"/> 
Figure 2. Diagram of a culture setup. The bead deposit is the source of proteins; the explants are placed to either side and all elements are covered with Matrigel.
<img src="/files/ftp_upload/2583/2583fig3.jpg" alt="Figure 3"/> 
Figure 3. Corresponding plots of cell tracks colored by quadrants (arrows point toward the bead deposit). The cell tracks on the right are migrating toward fraction D, which appears to attract cells; the cells on the left are migrating toward fraction A, which appears to repulse migrating cells. 
<img src="/files/ftp_upload/2583/2583fig4.jpg" alt="Figure 4"/> 
Figure 4. A) A series of images from selected timepoints showing a growing cloud of cells migrating out of an explant. An example cell, indicated by an arrow, is tracked through time as indicated below (scale bar 200&mu;m). This explant was visualized with live imaging for 38 h. Shown here are images of cells leaving the explant from the beginning of recording the images to the final image. B) The cell indicated in part A at the times indicated. The cell shows repetitive branching (scale bar 50&mu;m).

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Animal use:
All procedures involving animals were performed in accordance to the USUHS and NIH institutional guidelines and approved by the USUHS IACUC.

Here we present a simple system to study and analyze migratory behavior of neural cells in response to chemical cues. In vitro migration assays have been used to identify extracellular molecules affecting the locomotion of cells. Our paradigm ensures that each individual cell studied has a 3-dimensional (3D) environment with which to interact. It is a considerable advantage over cultures where cells migrating out of explants are forced to move on the flat surface of poly-D-lysine-coated coverslip, thus devoid of interactions with the extracellular matrix (e.g. Hirschberg et al. '10; 8. M&eacute;tin et al. '07). However with respect to the larger scale of tracking migrating cells, it remains 2-dimensional, as the cells are vertically constrained to the space between the glass coverslip on the bottom and the top surface of Matrigel, which is relatively thin compared to the horizontal freedom of migration. Such a setup allows for easier analysis of movement, which still remains essentially 2-dimensional. Similar 3D paradigms consisting of GE explants or re-aggregates of GE cells embedded in collagen or Matrigel, were previously employed, using aggregates of cells expressing a particular signaling protein as a source of chemical cues (e.g. Liodis et al. '07; Martini et al.. '09; N&oacute;brega-Pereira et al. '08; Wichterle et al. '03). The method presented here uses a slow release system in the form of latex beads with adhered proteins to allow for a sustained level in the local environment, which is higher than proteins added directly to the medium, making the system particularly suitable when the amount of available protein is limited or when a complex mixture of proteins is tested. The choice of beads is not limited to latex-based products, and includes also other materials, for example agarose beads, each with different characteristics. To further increase the density of the assay, we used two explants apposed to the beads deposit on opposite sides. To ensure a specific area of origin, we have used ganglionic eminence explants as a source of cells. We evaluated differences in medial and lateral portions of the ganglionic eminence, but did not see any differences. We should point out that this study uses ferrets; in this species the medial and lateral ganglionic eminences fuse earlier than in rodents (Poluch et al. '08). At the time the cultures were made (P0) the medial and lateral ganglionic eminences were fused, even though many cells populating the neocortex are still being generated and migrating. However, if a truly homogenous source of cells is preferred, a cell suspension can be prepared from the dissected tissue and directly mixed with the Matrigel, or alternatively spun down and a resulting pellet punched to make 'explants'. Use of enzymatic tissue digestion to prepare the cell suspension is discouraged, for even residual traces of proteolytic activity can liquefy Matrigel over the course of experiment. Also, a due diligence has to be applied to the timing of the culture setup. Although Matrigel requires a period of warming to polymerize, even a slightly extended period of exposure of small drops of Matrigel (as used in this protocol) to the air causes drying and results in a shell that is impenetrable to migrating cells and which cannot be erased by subsequent covering with fresh Matrigel.

<table border="1">
	<tbody> 
 <tr>
 	<th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</td>
 <th>Comments (optional)</th>	
 </tr>
 <tr>
 	<td>Neurobasal Medium (1X), liquid</td>
 <td>Invitrogen</td>
 <td>21103-049</td>
 <td>&nbsp;</td>	
 </tr>
 <tr>
 	<td>N-2 Supplement (100X), liquid</td>
 <td>Invitrogen</td>
 <td>17502-048</td>
 <td>&nbsp;</td>	
 </tr>
 <tr>
 	<td>B-27 Supplement (50X)</td>
 <td>Invitrogen</td>
 <td>0080085-SA</td>
 <td>&nbsp;</td>	
 </tr>
 <tr>
 	<td>BD Matrigel Basement Membrane Matrix</td>
 <td>BD Biosciences</td>
 <td>354234</td>
 <td>&nbsp;</td>	
 </tr>
 <tr>
 	<td>Harris Uni-Core, 0.5mm size</td>
 <td>Electron Microscopy Sciences</td>
 <td>69036-05</td>
 <td>Punch bore</td>	
 </tr>
 <tr>
 	<td>Green Retrobeads IX</td>
 <td>Lumafluor, Inc.</td>
 <td>&nbsp;</td>
 <td>Latex microspheres</td>	
 </tr>
 
 </tbody>
</table>

migratory behavior of cells generated in ganglionic eminence cultures

Migration of cells is a common process that leads to the development and maturation of the vertebrate central nervous system (Hatten, '99). The cerebral cortex consists of two basic neuronal types: excitatory and inhibitory. These cells arise in distinct areas and migrate into the cortex along different routes (Pearlman et al., '98). Inhibitory interneurons migrate tangentially from subcortical sources, mostly from different regions of the ganglionic eminences (Gelman et al., '09; Xu et al., '04). Their movement requires precise spatiotemporal control imposed by environmental cues, to allow for the establishment of proper cytoarchitecture and connectivity in the cerebral cortex (Caviness &amp; Rakic, '78; Hatten, '90; Rakic, '90). To study the migratory behavior of cells generated in proliferative zones of the ganglionic eminences (GE) in newborn ferrets in vitro we used a 3 dimensional culture arrangement in a BD Matrigel Matrix. The culture setup consisted of two GE explants and a source of tested proteins extracted from the cerebral cortex and adsorbed on fluorescent latex Retrobeads IX positioned between the explants (Hasling et al., '03; Riddle et al., '97). After 2-3 days of culture, the cells start to appear at the edge of the explant showing a propensity to leave the tissue in a radial direction. Live imaging allowed observation of migratory patterns without the necessity of labeling or marking the cells. When exposed to fractions of the protein extract obtained from isochronic ferret cortex, the GE cells displayed different behaviors as judged by quantitative kinetic analysis of individual moving cells.

Watch this Scientific Journal Video about Migratory Behavior of Cells Generated in Ganglionic Eminence Cultures at JoVE.com

Migratory Behavior of Cells Generated in Ganglionic Eminence Cultures

Time lapse imaging of 3D tissue culture allows studying migratory behavior of individual cells originating from ganglionic eminence in reaction to fractionated protein extract from cerebral cortex.

Migration of cells is a common process that leads to the development and maturation of the vertebrate central nervous system (Hatten, '99). The cerebral ...

migratory-behavior-of-cells-generated-in-ganglionic-eminence-cultures

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Neuroscience

9.9K Views.  Uniformed Services University. Временной интервал визуализации 3D культуры ткани позволяет изучать миграционное поведение отдельных клеток, происходящих из ганглиозные возвышение в качестве реакции на фракционированного извлечения белка из коры головного мозга.

Video: Migratory Behavior of Cells Generated in Ganglionic Eminence Cultures

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 ...

Cryopreservation of Cortical Tissue Blocks for the Generation of Highly Enriched Neuronal Cultures

In this study, we outline a standardized protocol for the successful cryopreservation and thawing of cortical brain tissue blocks to generate highly enriched neuronal cultures. For this protocol the freezing medium used is 10% dimethyl sulfoxide (DMSO) diluted in Hank's Buffered Salt Solution (HBSS). Blocks of cortical tissue are transferred to cryovials containing the freezing medium and slowly frozen at -1&deg;C/min in a rate-controlled freezing container. Post-thaw processing and dissociation of frozen tissue blocks consistently produced neuronal-enriched cultures which exhibited rapid neuritic growth during the first 5 days in culture and significant expansion of the neuronal network within 10 days. Immunocytochemical staining with the astrocytic marker glial fibrillary acidic protein (GFAP) and the neuronal marker beta-tubulin class III, revealed high numbers of neurons and astrocytes in the cultures. Generation of neural precursor cell cultures after tissue block dissociation resulted in rapidly expanding neurospheres, which produced large numbers of neurons and astrocytes under differentiating conditions. This simple cryopreservation protocol allows for the rapid, efficient, and inexpensive preservation of cortical brain tissue blocks, which grants increased flexibility for later generation of neuronal, astrocyte, and neuronal precursor cell cultures.

Here, we describe a method for efficient cryopreservation and thawing of cortical brain tissue blocks to generate highly enriched neuronal cultures. This simple protocol provides flexibility for later generation of neuronal, astrocyte, and neuronal precursor cell cultures.

In this study, we outline a standardized protocol for the successful cryopreservation and thawing of cortical brain tissue blocks to generate highly ...

Studying the Neural Basis of Adaptive Locomotor Behavior in Insects

Studying the neural basis of walking behavior, one often faces the problem that it is hard to separate the neuronally produced stepping output from those leg movements that result from passive forces and interactions with other legs through the common contact with the substrate. If we want to understand, which part of a given movement is produced by nervous system motor output, kinematic analysis of stepping movements, therefore, needs to be complemented with electrophysiological recordings of motor activity. The recording of neuronal or muscular activity in a behaving animal is often limited by the electrophysiological equipment which can constrain the animal in its ability to move with as many degrees of freedom as possible. This can either be avoided by using implantable electrodes and then having the animal move on a long tether (i.e. Clarac et al., 1987; Duch &amp; Pfl&uuml;ger, 1995; B&ouml;hm et al., 1997; Gruhn &amp; Rathmayer, 2002) or by transmitting the data using telemetric devices (Kutsch et al, 1993; Fischer et al., 1996; Tsuchida et al. 2004; Hama et al., 2007; Wang et al., 2008). Both of these elegant methods, which are successfully used in larger arthropods, often prove difficult to apply in smaller walking insects which either easily get entangled in the long tether or are hindered by the weight of the telemetric device and its batteries. In addition, in all these cases, it is still impossible to distinguish between the purely neuronal basis of locomotion and the effects exerted by mechanical coupling between the walking legs through the substrate. One solution for this problem is to conduct the experiments in a tethered animal that is free to walk in place and that is locally suspended, for example over a slippery surface, which effectively removes most ground contact mechanics. This has been used to study escape responses (Camhi and Nolen, 1981; Camhi and Levy, 1988), turning (Tryba and Ritzman, 2000a,b; Gruhn et al., 2009a), backward walking (Graham and Epstein, 1985) or changes in velocity (Gruhn et al., 2009b) and it allows the experimenter easily to combine intra- and extracellular physiology with kinematic analyses (Gruhn et al., 2006).
We use a slippery surface setup to investigate the timing of leg muscles in the behaving stick insect with respect to touch-down and lift-off under different behavioral paradigms such as straight forward and curved walking in intact and reduced preparations.

We describe a method to record motor activity, timed to the electrically recorded tarsal contact signal in a tethered insect, walking on a slippery surface. This is used to study the neural basis of adaptive behavior under reduced influence of mechanical interaction between legs through the substrate.

Studying the neural basis of walking behavior, one often faces the problem that it is hard to separate the neuronally produced stepping output from those ...

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

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 ...

Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling of auditory axons within the VIIIth cranial nerve would enhance the study of central auditory circuit development. This approach is challenging because multiple sensory organs of the inner ear contribute to the VIIIth nerve 1. Moreover, markers that reliably distinguish auditory versus vestibular groups of axons within the avian VIIIth nerve have yet to be identified. Auditory and vestibular pathways cannot be distinguished functionally in early embryos, as sensory-evoked responses are not present before the circuits are formed. Centrally projecting VIIIth nerve axons have been traced in some studies, but auditory axon labeling was accompanied by labeling from other VIIIth nerve components 2,3. Here, we describe a method for anterograde tracing from the acoustic ganglion to selectively label auditory axons within the developing VIIIth nerve. First, after partial dissection of the anterior cephalic region of an 8-day chick embryo immersed in oxygenated artificial cerebrospinal fluid, the cochlear duct is identified by anatomical landmarks. Next, a fine pulled glass micropipette is positioned to inject a small amount of rhodamine dextran amine into the duct and adjacent deep region where the acoustic ganglion cells are located. Within thirty minutes following the injection, auditory axons are traced centrally into the hindbrain and can later be visualized following histologic preparation. This method provides a useful tool for developmental studies of peripheral to central auditory circuit formation.

Here we describe a microdissection technique followed by fluorescent dye injection into the acoustic ganglion of early chick embryos for selective tracing of auditory axon fibers in the nerve and hindbrain.

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Assays to Detect UV-reflecting Structures and Determine their Importance in Mate Preference using the Sailfin Molly Poecilia latipinna

Many organisms use cues and signals beyond human sensitivity during social interactions. It is important to take into account how organisms perceive their worlds when trying to understand their behavior and ecology. Sensitivity to the ultraviolet spectrum (UV; 300 - 400 nm) is found across multiple genera of birds, fish, reptiles, amphibians, and even mammals. This protocol describes a technique for examining organisms for the presence of UV-reflecting structures and a method for testing whether these cues are used as social signals in the context of mate choice. A spectrophotometer is used to detect the presence of UV reflectance and variation in reflective intensity between individuals and sexes. An example of this technique is presented in which a dichotomous mate choice test exposes sexually receptive individuals to opposite sex individuals whose visual appearance can be manipulated by filters that either transmit full spectrum or block UV wavelengths. This system allowed for the determination that female, but not male, sailfin mollies (Poecilia latipinna) were using UV markings as part of their mating decisions. These types of studies serve to expand our knowledge of the range of organisms that utilize UV and provide insight into how UV plays a role in their lives.

Assays to Detect UV-reflecting Structures and Determine their Importance in Mate Preference using the Sailfin Molly Poecilia latipinna

This protocol outlines the use of spectrophotometry to detect ultraviolet-reflecting structures on organisms (in this example, the sailfin molly Poecilia latipinna) and describes dichotomous choice tests for fish that allow inferences to be made on the role of ultraviolet cues during mate selection.

Many organisms use cues and signals beyond human sensitivity during social interactions. It is important to take into account how organisms perceive their ...

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.

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

Combining Behavior and EEG to Study the Effects of Mindfulness Meditation on Episodic Memory

Although there has been recent interest in how mindfulness meditation can affect episodic memory as well as brain structure and function, no study has examined the behavioral and neural effects of mindfulness meditation on episodic memory. Here we present a protocol that combines mindfulness meditation training, an episodic memory task, and EEG to examine how mindfulness meditation changes behavioral performance and the neural correlates of episodic memory. Subjects in a mindfulness meditation experimental group were compared to a waitlist control group. Subjects in the mindfulness meditation experimental group spent four weeks training and practicing mindfulness meditation. Mindfulness was measured before and after training using the Five Facet Mindfulness Questionnaire (FFMQ). Episodic memory was measured before and after training using a source recognition task. During the retrieval phase of the source recognition task, EEG was recorded. The results showed that mindfulness, source recognition behavioral performance, and EEG theta power in right frontal and left parietal channels increased following mindfulness meditation training. In addition, increases in mindfulness correlated with increases in theta power in right frontal channels. Therefore, results obtained from combining mindfulness meditation training, an episodic memory task, and EEG reveal the behavioral and neural effects of mindfulness meditation on episodic memory.

Here we present a protocol for combining mindfulness meditation training, an episodic memory task, and EEG to understand the behavioral and neural effects of mindfulness meditation on episodic memory.

Although there has been recent interest in how mindfulness meditation can affect episodic memory as well as brain structure and function, no study has ...