This work was supported by Canadian Institutes of Health Research (CIHR) Grant to Wayne S. Sossin. Carole A. Farah is the recipient of a postdoctoral fellowship from the Fonds de la Recherche en Sant&eacute; du Qu&eacute;bec (FRSQ) and a Conrad Harrington fellowship. The authors would like to thank Joanna Bougie, Margaret Hastings and Margaret Labban for help with the video shoot and Madeline Richmond Pool for help with the graphic overview.

1. Preparation and Maintenance of Sf9 Cell Monolayer Cultures 
<ol>
 <li>Sf9 cell culture and maintenance is performed in a tissue culture hood.</li>
 <li>Sf9 cells are derived from Spodoptera frugiperda ovarian cells (Sf21 cells) and can be purchased as frozen cells in Grace's media from Invitrogen.</li>
 <li>Place 8 mL of Grace's insect medium, supplemented (1X), to which 30% fetal bovine serum (FBS) has been added in a 75 cm2 cell culture flask with a canted neck.</li>
 <li>Thaw the frozen tube of Sf9 cells rapidly in a 37&deg;C water bath by moving the tube back and forth (about 1-2 minutes).</li>
 <li>Transfer Sf9 cells to the 75 cm2 cell culture flask and rock plate gently by hand to distribute the cells evenly.</li>
 <li>Incubate 1-2 hours at 27&deg;C until cells have attached.</li>
 <li>Remove old medium and replace with 10 mL of freshGrace's insect medium with 30% FBS.</li>
 <li>Incubate at 27&deg;C until the cells are confluent.</li>
 <li>To maintain monolayer cultures, remove old medium from the confluent flask of Sf9 cells and add 5 mL of Grace's insect medium with 10% FBS.</li>
 <li>Lightly tap on the side of the flask several times to detach Sf9 cells.</li>
 <li>Using a 10 mL pipette and a pipettor, pipette the media up and down once gently, spraying flask wall while pipetting down. Transfer the cells to a sterile 15 mL Polysterene tube.</li>
 <li>Add 2 mL of Sf9 cell suspension to 8 mL of Grace's insect medium with 10% FBS (1:5 dilution to maintain log phase growth) in new 75 cm2 cell culture flask and rock plate gently by hand.</li>
 <li>Incubate at 27&deg;C until the cells are confluent.</li>
</ol>
2. Expression of Fluorescently Tagged PKCs in Sf9 Cells
<ol>
 <li>For live-imaging experiment, culture Sf9 cells on 35 mm MatTek glass bottom culture disheswith a glass surface of 14 mm and a coverslip thickness of 0.16 to 0.19 mm.</li>
 <li>PKCs tagged with fluorescent proteins are expressed in Sf9 cells using Cellfectin II transfection reagent following recommendation of the manufacturer with the modifications detailed below.</li>
 <li>Day 1: Prior to plating the cells, treat each MatTek dish with Grace's insect medium with 10% FBS for at least 30 minutes.</li>
 <li>Count Sf9 cells from step 11 (above) using a hemacytometer.</li>
 <li>Plate 0.05 x 106 cells on each MatTek glass coverslip in a total volume of 150 &mu;L. Allow the cells to attach for 30 minutes at room temperature.</li>
 <li>Add 2 mL of Grace's supplemented with 10% FBS to each dish 1 mL at a time slowly to avoid detaching the cells.</li>
 <li>Incubate at 27&deg;C overnight.</li>
 <li>From this point on, use manufacturer's recommendation for transfection of Sf9 cells with plasmid DNA.</li>
 <li>Day 2: Transfect Sf9 cells with fluorescently tagged PKCs using Cellfectin II transfection reagent. We have often co-expressed in this system PKCs tagged with enhanced Green Fluorescent Protein (eGFP) and monomeric Red Fluorescent Protein (mRFP) (details for plasmid construction have been previously described 2,5). Expression of one fluorescent protein at a time yields about 70-80% expressing cells 72 hours post-transfection. Co-expression of two fluorescent proteins decreases transfection efficiency to about 30% co-expressing cells. When co-expressing eGFP-tagged PKC and mRFP-tagged PKC, use 2x more plasmid DNA for the mRFP-tagged protein for optimal protein expression.</li>
 <li>Perform live-imaging 48-72 hours post-transfection (for optimal protein expression, wait 72 hours).</li>
</ol>
3. Expression of Fluorescently Tagged PKCs in Aplysia Sensory Neurons
<ol>
 <li>Preparation of Aplysia neuronal cell cultures has been described in a video article by Zhao and colleagues 4.</li>
 <li>PKCs tagged with fluorescent proteins are expressed in Aplysia sensory neurons using microinjection procedure detailed below.</li>
 <li>Prepare a stock solution of 1% Fast Green (FCF) in distilled water, filter using a syringe and a 28 mm syringe filter (0.20 &mu;m). Aliquot and store at -20&deg;C.&nbsp;</li>
 <li>Microinjection of the sensory neurons can be performed on day 1 or day 2 after isolating the neurons but we find that waiting 2 days allows for better cell adherence to the coverslip.</li>
 <li>On day 2 after isolation of the sensory neurons, thaw an aliquot of Fast Green and filter it again using a syringe and a 28 mm syringe filter (0.20 &mu;m).</li>
 <li>Prepare a solution of plasmid DNA in distilled water containing 0.5% Fast Green. Concentrations of plasmid DNA as high as 0.4 &mu;g/&mu;L can be used but it is not recommended to go above these values.&nbsp;</li>
 <li>Filter the plasmid DNA / Fast Green solution using a 1 mL syringe and a 4 mm syringe filter (0.20 &mu;m).</li>
 <li>Centrifuge the plasmid DNA / Fast Green solution at 16110 x g for 15 minutes using a microcentrifuge.</li>
 <li>Prepare sharp electrodes for microinjection of neurons using a microelectrode puller (Sutter Flaming/Brown Micropipette Puller, Model P-97). Using a box filament, pull glass microelectrodes with 1 mm outer diameter, 0.75 mm inside diameter and 100 mm length using thin wall glass capillaries with a filament inside (World Precision Instruments TW100F-4). For more information about pulling microinjection electrodes please refer to Sutter electrode cookbook.</li>
 <li>Fill each electrode with 2 &mu;L of plasmid DNA / Fast Green solution using microloader pipette tip (eppendorf CA32950-050).</li>
 <li>Transfer the MatTek glass bottom culture dish containing Aplysia neuronal cultures to the microinjection station.</li>
 <li>The microinjection station consists of the following: a home-made large glass stage to hold the culture dishes on a vibration isolation table (Kinetic Systems vibration isolation workstation), a stereo-scope with external halogen illumination, a micromanipulator (Sutter Huxley Wall Type Micromanipulator which allows both coarse and ultra-fine positioning in the three-axis), a microelectrode holder connected to a pneumatic pump (World Precision Instruments PV820 Pneumatic Pico-Pump). The pressure Input of the pneumatic pump is connected to a compressed Nitrogen tank.&nbsp;&nbsp;&nbsp;</li>
 <li>Insert a microelectrode into the microelectrode holder.</li>
 <li>Viewing under a stereo-microscope, insert the tip of the micropipette into the cell nucleus.</li>
 <li>Deliver short pressure pulses of Nitrogen (10 to 100 ms in duration; 20 lb/in2) until the nucleus becomes uniformely green.</li>
 <li>Incubate the cells for 4 hours at room temperature and keep at 4&deg;C until use. For optimal PKC translocation, perform live-imaging experiment 20 - 24 hours post-injection.</li>
</ol>
4. Visualization of PKC Translocation in Living Sf9 Cells and in Aplysia Sensory Neurons
<ol>
 <li>For imaging, we use a Zeiss LSM510 inverted confocal microscope with an Axiovert 200.</li>
 <li>Imaging protocol is the same for Sf9 cells and for Aplysia sensory neurons except for the following: Sf9 cells are imaged with a x63 oil immersion objective whereas Aplysia sensory neurons are imaged with a x40 oil immersion objective. Sf9 cells are imaged in a temperature-controlled chamber maintained at 26-27oC whereas Aplysia sensory neurons are imaged at room temperature maintained around 18-20oC.</li>
 <li>We use a 30 mW Argon laser (excitation at 488nm) with 50% laser output to image PKC tagged with eGFP protein and a HeNe laser (excitation at 543nm) to image PKC tagged with mRFP protein. The Argon and HeNe laser lines are attenuated to 4% and 50% transmission output respectively prior to imaging and the pinhole is adjusted to one.</li>
 <li>It is important to have the culture dish stabilized on the imaging stage and to avoid movement or vibration.</li>
 <li>Remove 1 mL of culture media from the dish gently using a 10 mL pipette leaving 1 mL total volume in the dish.</li>
 <li>Take pictures in the middle section of the cells where the nucleus can be seen.</li>
 <li>Set up a time series of 10-20 confocal images taking a picture every 30 seconds.</li>
 <li>Start the time series.</li>
 <li>After 2 pictures are taken (after the 30 seconds time point), add 1 mL of the drug (2X concentration) gently drop-by-drop on top of the cells using a P1000 pipette. The drug is added continuously for about 30 seconds.</li>
 <li>Some drugs induce a fast translocation which can be visible immediately following drug treatment (at the 60 seconds time point) while others induce a slow translocation which is only optimal 5-10 minutes post-treatment (Movie 1 and Movie 2).</li>
 <li>If the drug needs to be washed away, use 2 x 10 mL pipets to do the wash, pipetting the wash buffer down gently with one pipette on one side of the dish and pipeting up the media on the other side.</li>
 <li>If the wash has been effective and if the effect of the drug is reversible, PKC reverts back to the cytosol within 30-60 seconds.</li>
 <li>Save the movie as an LSM file.</li>
 <li>Use NIH Image J software to open the LSM files and quantify the images Pre and Post drug treatment. Quantification has been described elsewhere 1.</li>
</ol>

<ol>
	<li>Farah, C. A., Nagakura, I., Weatherill, D., Fan, X. &. a. m. p. ;. a. m. p., Sossin, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiological+role+for+phosphatidic+acid+in+the+translocation+of+the+novel+protein+kinase+C+Apl+II+in+Aplysia+neurons.">Physiological role for phosphatidic acid in the translocation of the novel protein kinase C Apl II in Aplysia neurons.</a> Mol Cell Biol. 28, 4719-4733 (2008).</li><li>Zhao, Y., Leal, K., Abi-Farah, C., Martin, K. C., Sossin, W. S. &. a. m. p. ;. a. m. p., Klein, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=M.Isoform+specificity+of+PKC+translocation+in+living+Aplysia+sensory+neurons+and+a+role+for+Ca2+-dependent+PKC+APL+I+in+the+induction+of+intermediate-term+facilitation.">M.Isoform specificity of PKC translocation in living Aplysia sensory neurons and a role for Ca2+-dependent PKC APL I in the induction of intermediate-term facilitation.</a> J Neurosci. 26, 8847-8856 (2006).</li><li>Farah, C. A., Weatherill, D., Dunn, T. W. &. a. m. p. ;. a. m. p., Sossin, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PKC+differentially+translocates+during+spaced+and+massed+training+in+Aplysia.">PKC differentially translocates during spaced and massed training in Aplysia.</a> J Neurosci. 29, 10281-10286 (2009).</li><li>Zhao, Y., Wang, D. O., Martin, K. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+Aplysia+sensory-motor+neuronal+cell+cultures.">Preparation of Aplysia sensory-motor neuronal cell cultures.</a> J Vis Exp. , (2009).</li><li>Manseau, F., Fan, X., Hueftlein, T., Sossin, W. S., Castellucci, V. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ca2+-independent+PKC+Apl+II+mediates+the+serotonin+induced+facilitation+at+depressed+synapses+in+Aplysia.">Ca2+-independent PKC Apl II mediates the serotonin induced facilitation at depressed synapses in Aplysia.</a> J. Neuroscience. 21, 1247-1256 (2001).</li></ol>

No conflicts of interest declared.

We have described a technique to image translocation of fluorescently tagged PKCs in real time in Sf9 cells and in Aplysia sensory neurons. Sf9 cells provide a simple system to image PKC translocation since culturing and transfecting them is pretty straight-forward. In contrast, microinjection of Aplysia sensory neurons takes some time to master; up to a few months. Difficulties related to this technique include electrode clogging. Filtering the injection solution and centrifuging it usually helps but s...

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Sf9 frozen cells</td>
<td>Spodoptera frugiperda ovarian cells</td>
<td>Invitrogen</td>
<td>B825-01</td>
<td>&nbsp;</td>
<tr>
<td>Grace&#x2019;s Insect Medium, Supplemented (1X), liquid</td>
<td>Reagent</td>
<td>Gibco</td>
<td>11605-094</td>
<td>Use to culture Sf9 cells</td>
</tr>
<tr>
<td>Corning 75cm2 canted neck cell culture flask</td>
<td>Tool</td>
<td>Corning</td>
<td>430720</td>
<td>Use to culture Sf9 cells</td>
</tr>
<tr>
<td>Fetal Bovine Serum</td>
<td>Reagent</td>
<td>CanSera</td>
<td>CS-C08-500</td>
<td>Supplement Grace's media with FBS prior to use</td>
</tr>
<tr>
<td>Syringe</td>
<td>Tool</td>
<td>Becton Dickinson</td>
<td>309604</td>
<td>Use to filter Fast Green solution</td>
</tr>
<tr>
<td>Syringe</td>
<td>Tool</td>
<td>Becton Dickinson</td>
<td>309602</td>
<td>Use to filter plasmid DNA / Fast Green solution</td>
</tr>
<tr>
<td>Filter</td>
<td>Tool</td>
<td>Corning</td>
<td>431229</td>
<td>Use to filter Fast Green solution</td>
</tr>
<tr>
<td>Filter</td>
<td>Tool</td>
<td>Corning</td>
<td>431212</td>
<td>Use to filter plasmid DNA / Fast Green solution</td>
</tr>
<tr>
<td>Glass Bottom Dish</td>
<td>Tool</td>
<td>Mattek</td>
<td>P35G-1.5-14-C</td>
<td>Use to culture Sf9 cells prior to transfection and imaging</td>
</tr>
<tr>
<td>Cellfectin II Reagent</td>
<td>Reagent</td>
<td>Invitrogen</td>
<td>10362-100</td>
<td>Used to express fluorescently tagged PKCs in Sf9 cells</td>
</tr>
<tr>
<td>Fast Green FCF</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>F-7252</td>
<td>Use to visualize microinjection of plasmid DNA solution into Aplysia sensory neurons</td>
</tr>
<tr>
<td>Thin wall glass capillaries</td>
<td>Tool</td>
<td>World Precision Instruments</td>
<td>TW100F-4</td>
<td>Use to make microelectrodes for microinjection</td>
</tr>
<tr>
<td>Microloader pipette tip</td>
<td>Tool</td>
<td>Eppendorf</td>
<td>CA32950-050</td>
<td>Autoclave before using to fill up the microelectrodes for microinjection</td>
</tr>
<tr>
<td>PV820 Pneumatic PicoPump </td>
<td>Tool</td>
<td>World Precision Instruments</td>
<td>SYS-PV820</td>
<td>Part of microinjection station</td>
</tr>
<tr>
<td>Microelectrode holder</td>
<td>Tool</td>
<td>World Precision Instruments</td>
<td>MPH6S</td>
<td>Use to hold microelectrode </td>
</tr>
<tr>
<td>Micromanipulator</td>
<td>Tool</td>
<td>Sutter</td>
<td>MP-85</td>
<td>Use to position microelectrode in the three-axis</td>
</tr>		</tbody>
		</table>
		</div>

live-imaging of pkc translocation in sf9 cells and in aplysia sensory neurons

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and learning and memory. There are four known families of PKC isoforms in vertebrates: classical PKCs (&alpha;, &beta;I, &beta;II and &gamma;), novel type I PKCs (&#949; and &eta;), novel type II PKCs (&delta; and &theta;), and atypical PKCs (&zeta; and &iota;). The classical PKCs are activated by Ca2+ and diacylclycerol (DAG), while the novel PKCs are activated by DAG, but are Ca2+-independent. The atypical PKCs are activated by neither Ca2+ nor DAG. In Aplysia californica, our model system to study memory formation, there are three nervous system specific PKC isoforms one from each major class, namely the conventional PKC Apl I, the novel type I PKC Apl II and the atypical PKC Apl III. PKCs are lipid-activated kinases and thus activation of classical and novel PKCs in response to extracellular signals has been frequently correlated with PKC translocation from the cytoplasm to the plasma membrane. Therefore, visualizing PKC translocation in real time in live cells has become an invaluable tool for elucidating the signal transduction pathways that lead to PKC activation. For instance, this technique has allowed for us to establish that different isoforms of PKC translocate under different conditions to mediate distinct types of synaptic plasticity and that serotonin (5HT) activation of PKC Apl II requires production of both DAG and phosphatidic acid (PA) for translocation 1-2. Importantly, the ability to visualize the same neuron repeatedly has allowed us, for example, to measure desensitization of the PKC response in exquisite detail 3. In this video, we demonstrate each step of preparing Sf9 cell cultures, cultures of Aplysia sensory neurons have been described in another video article 4, expressing fluorescently tagged PKCs in Sf9 cells and in Aplysia sensory neurons and live-imaging of PKC translocation in response to different activators using laser-scanning microscopy.

Watch this Scientific Journal Video about Live-imaging of PKC Translocation in Sf9 Cells and in Aplysia Sensory Neurons at JoVE.com

Live-imaging of PKC Translocation in Sf9 Cells and in Aplysia Sensory Neurons

In this video, we demonstrate visualization of PKC translocation in living cells using fluorescently tagged PKCs.

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and ...

live-imaging-pkc-translocation-sf9-cells-aplysia-sensory

Research

JoVE Journal

Neuroscience

11.9K vues.  McGill University. Dans cette vidéo, nous démontrons la visualisation de la translocation de la PKC dans les cellules vivantes en utilisant PKC fluorescence marqués.

Vidéo: Live-imaging of PKC Translocation in Sf9 Cells and in Aplysia Sensory Neurons

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

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 ...

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the olfactory bulb (OB). The molecular mechanisms of OSN axon growth and targeting are not well understood. Manipulating gene expression and subsequent 
visualizing of single OSN axons and their terminal arbor morphology have thus far been challenging. To study gene function at the single cell level within a specified 
time frame, we developed a lentiviral based technique to manipulate gene expression in OSNs in vivo. Lentiviral particles are delivered to OSNs by microinjection 
into the olfactory epithelium (OE). Expression cassettes are then permanently integrated into the genome of transduced OSNs. Green fluorescent protein expression 
identifies infected OSNs and outlines their entire morphology, including the axon terminal arbor. Due to the short turnaround time between microinjection and 
reporter detection, gene function studies can be focused within a very narrow period of development. With this method, we have detected GFP expression within as 
few as three days and as long as three months following injection. We have achieved both over-expression and shRNA mediated knock-down by lentiviral microinjection. 
This method provides detailed morphologies of OSN cell bodies and axons at the single cell level in vivo, and thus allows characterization of candidate gene function 
during olfactory development.

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

We present a lentiviral technique for genetic manipulation and visualization of single olfactory sensory neuron axon and its terminal arborization in vivo.

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the ...

Flash Photolysis of Caged Compounds in the Cilia of Olfactory Sensory  Neurons

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. Caged compounds are molecules made physiologically inactive by a chemical cage that can be broken by a flash of ultraviolet light. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged compounds for the study of olfactory transduction in dissociated mouse olfactory sensory neurons. The process of olfactory transduction (Figure 1) takes place in the cilia of olfactory sensory neurons, where odorant binding to receptors leads to the increase of cAMP that opens cyclic nucleotide-gated (CNG) channels2. Ca entry through CNG channels activates Ca-activated Cl channels. We show how to dissociate neurons from the mouse olfactory epithelium3 and how to activate CNG channels or Ca-activated Cl channels by photolysis of caged cAMP4 or caged Ca5. We use a flash lamp6,7 to apply ultraviolet flashes to the ciliary region to uncage cAMP or Ca while patch-clamp recordings are taken to measure the current in the whole-cell voltage-clamp configuration8-11.

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged cAMP or caged Ca for the study of olfactory transduction in dissociated mouse olfactory sensory neurons.

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. ...

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.
This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath5, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.
To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation4,6,7. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.
We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g. in the suspended buccal mass preparation8 or in vivo9. This process can also be applied in other motor pools10,11,12 in Aplysia or in other animal systems2,3,13,14.

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

In animals with large  identified neurons (e.g. mollusks), analysis of motor pools is done using  intracellular techniques1,2,3,4. Recently,  we developed ...

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

The marine gastropod mollusk Aplysia californica has a venerable history as a model of nervous system function, with particular significance in studies of learning and memory. The typical preparations for such studies are ones in which the sensory and motoneurons are left intact in a minimally dissected animal, or a technically elaborate neuronal co-culture of individual sensory and motoneurons. Less common is the isolated neuronal preparation in which small clusters of nominally homogeneous neurons are dissociated into single cells in short term culture. Such isolated cells are useful for the biophysical characterization of ion currents using patch clamp techniques, and targeted modulation of these conductances. A protocol for preparing such cultures is described. The protocol takes advantage of the easily identifiable glutamatergic sensory neurons of the pleural and buccal ganglia, and describes their dissociation and minimal maintenance in culture for several days without serum.

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

We describe the dissection of the nervous system of the marine sea hare Aplysia after anesthesia, the isolation of neurons for short term-tissue culture, and recordings of single cell ion currents via the patch clamp technique.

The marine gastropod mollusk Aplysia californica has a venerable  history as a model of nervous system function, with particular significance in  studies ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

A Method to Target and Isolate Airway-innervating Sensory Neurons in Mice

Somatosensory nerves transduce thermal, mechanical, chemical, and noxious stimuli caused by both endogenous and environmental agents. The cell bodies of these afferent neurons are located within the sensory ganglia. Sensory ganglia innervate a specific organ or portion of the body. For instance, the dorsal root ganglia (DRG) are located in the vertebral column and extend processes throughout the body and limbs. The trigeminal ganglia are located in the skull and innervate the face, and upper airways. Vagal afferents of the nodose ganglia extend throughout the gut, heart, and lungs. The nodose neurons control a diverse array of functions such as: respiratory rate, airway irritation, and cough reflexes. Thus, to understand and manipulate their function, it is critical to identify and isolate airway specific neuronal sub-populations. In the mouse, the airways are exposed to a fluorescent tracer dye, Fast Blue, for retrograde tracing of airway-specific nodose neurons. The nodose ganglia are dissociated and fluorescence activated cell (FAC) sorting is used to collect dye positive cells. Next, high quality ribonucleic acid (RNA) is extracted from dye positive cells for next generation sequencing. Using this method airway specific neuronal gene expression is determined.

Organ specific sensory neurons are difficult to identify. Fast Blue tracing is used to identify nodose neurons innervating the airways for cell sorting. Sorted nodose neurons are used to extract high quality ribonucleic acid (RNA) for sequencing. Using this protocol, gene expression of airway specific neurons is determined.

Somatosensory nerves transduce thermal, mechanical, chemical, and noxious stimuli caused by both endogenous and environmental agents. The cell bodies of ...