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...

<table><tbody><tr><td>Sf9 frozen cells</td><td>&lt;em&gt;Spodoptera frugiperda&lt;/em&gt; ovarian cells</td><td>Invitrogen</td><td>B825-01</td><td></td></tr><tr><td>Grace&amp;rsquo;s Insect Medium, Supplemented (1X), liquid</td><td>Reagent</td><td>GIBCO, by Life Technologies</td><td>11605-094</td><td>Use to culture Sf9 cells</td></tr><tr><td>Corning 75cm&lt;sup&gt;2&lt;/sup&gt; 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&amp;rsquo;s media with FBS prior to use</td></tr><tr><td>Syringe</td><td>Tool</td><td>BD Biosciences</td><td>309604</td><td>Use to filter Fast Green solution</td></tr><tr><td>Syringe</td><td>Tool</td><td>BD Biosciences</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 Corp.</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 &lt;em&gt;Aplysia&lt;/em&gt; sensory neurons</td></tr><tr><td>Thin wall glass capillaries</td><td>Tool</td><td>World Precision Instruments, Inc.</td><td>TW100F-4</td><td>Use to make micr&amp;#8211;lectrodes 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 micr&amp;#8211;lectrodes for microinjection</td></tr><tr><td>PV820 Pneumatic PicoPump </td><td>Tool</td><td>World Precision Instruments, Inc.</td><td>SYS-PV820</td><td>Part of microinjection station</td></tr><tr><td>Micr&amp;#8211;lectrode holder</td><td>Tool</td><td>World Precision Instruments, Inc.</td><td>MPH6S</td><td>Use to hold micr&amp;#8211;lectrode </td></tr><tr><td>Micromanipulator</td><td>Tool</td><td>Sutter Instrument Co.</td><td>MP-85</td><td>Use to position micr&amp;#8211;lectrode in the three-axis</td></tr></tbody></table>

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 Views.  McGill University. In deze video laten we zien visualisatie van PKC translocatie in levende cellen met behulp van fluorescent gelabelde PKCS.

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

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 ...

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

Bioengineering

Monitoring Changes in the Intracellular Calcium Concentration and Synaptic Efficacy in the Mollusc Aplysia

It has been suggested that changes in intracellular calcium mediate the induction of a number of important forms of synaptic plasticity (e.g., homosynaptic facilitation) 1. These hypotheses can be tested by simultaneously monitoring changes in intracellular calcium and alterations in synaptic efficacy. We demonstrate how this can be accomplished by combining calcium imaging with intracellular recording techniques. Our experiments are conducted in a buccal ganglion of the mollusc Aplysia californica. This preparation has a number of experimentally advantageous features: Ganglia can be easily removed from Aplysia and experiments use adult neurons that make normal synaptic connections and have a normal ion channel distribution. Due to the low metabolic rate of the animal and the relatively low temperatures (14-16 &deg;C) that are natural for Aplysia, preparations are stable for long periods of time. 
To detect changes in intracellular free calcium we will use the cell impermeant version of Calcium Orange 2 which is easily 'loaded' into a neuron via iontophoresis. When this long wavelength fluorescent dye binds to calcium, fluorescence intensity increases. Calcium Orange has fast kinetic properties 3 and, unlike ratiometric dyes (e.g., Fura 2), requires no filter wheel for imaging. It is fairly photo stable and less phototoxic than other dyes (e.g., fluo-3) 2,4. Like all non-ratiometric dyes, Calcium Orange indicates relative changes in calcium concentration. But, because it is not possible to account for changes in dye concentration due to loading and diffusion, it can not be calibrated to provide absolute calcium concentrations. 
An upright, fixed stage, compound microscope was used to image neurons with a CCD camera capable of recording around 30 frames per second. In Aplysia this temporal resolution is more than adequate to detect even a single spike induced alteration in the intracellular calcium concentration. Sharp electrodes are simultaneously used to induce and record synaptic transmission in identified pre- and postsynaptic neurons. At the conclusion of each trial, a custom script combines electrophysiology and imaging data. To ensure proper synchronization we use a light pulse from a LED mounted in the camera port of the microscope. Manipulation of presynaptic calcium levels (e.g. via intracellular EGTA injection) allows us to test specific hypotheses, concerning the role of intracellular calcium in mediating various forms of plasticity.

Monitoring Changes in the Intracellular Calcium Concentration and Synaptic Efficacy in the Mollusc Aplysia

We demonstrate how changes in the intracellular free calcium concentration and synaptic efficacy can be simultaneously monitored in a ganglion preparation of Aplysia. We image intracellular calcium using a fluorescent dye, Calcium Orange, and induce and monitor synaptic transmission with sharp (intracellular) electrodes.

It has been suggested that changes in intracellular calcium mediate the induction of a number of important forms of synaptic plasticity (e.g., ...

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Biology

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of the fly such as imaginal discs, the larval brain or egg chambers of adult females or the adult intestine can be extracted and kept in culture for imaging with time-lapse microscopy, providing valuable insights into cell and developmental biology. Here, we describe in detail our current protocol for the dissection of Drosophila larval brains, and then present our current approach for immobilizing and orienting larval brains and other tissues on a glass coverslip using Fibrin clots. This immobilization method only requires the addition of Fibrinogen and Thrombin to the culture medium. It is suitable for high-resolution time lapse imaging on inverted microscopes of multiple samples in the same culture dish, minimizes the lateral drifting frequently caused by movements of the microscope stage in multi-point visiting microscopy and allows for the addition and removal of reagents during the course of imaging. We also present custom-made macros that we routinely use to correct for drifting and to extract and process specific quantitative information from time-lapse analysis.

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

This protocol describes applications of sample immobilization using Fibrin clots, limiting drifting, and allowing the addition and washout of reagents during live imaging. Samples are transferred to a drop of Fibrinogen containing culture medium on the surface of a coverslip, after which polymerization is induced by adding thrombin.

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of ...

Developmental Biology

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

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

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

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

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

Visualizing Protein Kinase A Activity In Head-fixed Behaving Mice Using In Vivo Two-photon Fluorescence Lifetime Imaging Microscopy

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. Despite their importance, technologies for tracking neuromodulatory events with cellular resolution are just beginning to emerge. Neuromodulators, such as dopamine, norepinephrine, acetylcholine, and serotonin, trigger intracellular signaling events via their respective G protein-coupled receptors to modulate neuronal excitability, synaptic communications, and other neuronal functions, thereby regulating information processing in the neuronal network. The above mentioned neuromodulators converge onto the cAMP/protein kinase A (PKA) pathway. Therefore, in vivo PKA imaging with single-cell resolution was developed as a readout for neuromodulatory events in a manner analogous to calcium imaging for neuronal electrical activities. Herein, a method is presented to visualize PKA activity at the level of individual neurons in the cortex of head-fixed behaving mice. To do so, an improved A-kinase activity reporter (AKAR), called tAKAR&#945;, is used, which is based on F&#246;rster resonance energy transfer (FRET). This genetically-encoded PKA sensor is introduced into the motor cortex via in utero electroporation (IUE) of DNA plasmids, or stereotaxic injection of adeno-associated virus (AAV). FRET changes are imaged using two-photon fluorescence lifetime imaging microscopy (2pFLIM), which offers advantages over ratiometric FRET measurements for quantifying FRET signal in light-scattering brain tissue. To study PKA activities during enforced locomotion, tAKAR&#945; is imaged through a chronic cranial window above the cortex of awake, head-fixed mice, which run or rest on a speed-controlled motorized treadmill. This imaging approach will be applicable to many other brain regions to study corresponding behavior-induced PKA activities and to other FLIM-based sensors for in vivo imaging.

A procedure is presented to visualize protein kinase A activities in head-fixed, behaving mice. An improved A-kinase activity reporter, tAKAR&#945;, is expressed in cortical neurons and made accessible for imaging through a cranial window. Two-photon fluorescence lifetime imaging microscopy is used to visualize PKA activities in vivo during enforced locomotion.

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. ...

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the cytoplasm or the release of intracellular Ca2+ stores. Pirt-GCaMP3-based Ca2+ imaging of primary sensory neurons of the dorsal root ganglion (DRG) in mice offers the advantage of simultaneous measurement of a large number of cells. Up to 1,800 neurons can be monitored, allowing neuronal networks and somatosensory processes to be studied as an ensemble in their normal physiological context at a populational level in vivo. The large number of neurons monitored allows the detection of activity patterns that would be challenging to detect using other methods. Stimuli can be applied to the mouse hindpaw, allowing the direct effects of stimuli on the DRG neuron ensemble to be studied. The number of neurons producing Ca2+ transients as well as the amplitude of Ca2+ transients indicates sensitivity to specific sensory modalities. The diameter of neurons provides evidence of activated fiber types (non-noxious mechano vs. noxious pain fibers, A&#946;, A&#948;, and C fibers). Neurons expressing specific receptors can be genetically labeled with td-Tomato and specific Cre recombinases together with Pirt-GCaMP. Therefore, Pirt-GCaMP3 Ca2+ imaging of DRG provides a powerful tool and model for the analysis of specific sensory modalities and neuron subtypes acting as an ensemble at the populational level to study pain, itch, touch, and other somatosensory signals.

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

This protocol describes the surgical exposure of the dorsal root ganglion (DRG) followed by GCaMP3 (genetically-encoded Ca2+ indicator; Green Fluorescent Protein-Calmodulin-M13 Protein 3) Ca2+ imaging of the neuronal ensembles using Pirt-GCaMP3 mice while applying a variety of stimuli to the ipsilateral hind paw.

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the ...

Subcellular Imaging of Neuronal Calcium Handling In Vivo

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a crucial component of intracellular signaling. The visualization of subcellular Ca2+ dynamics in vivo,&#160;where neurons can be studied in their native, intact circuitry, has proven technically challenging in complex nervous systems. The transparency and relatively simple nervous system of the nematode Caenorhabditis elegans enable the cell-specific expression and in vivo visualization of fluorescent tags and indicators. Among these are fluorescent indicators that have been modified for use in the cytoplasm as well as various subcellular compartments, such as the mitochondria. This protocol enables non-ratiometric Ca2+ imaging in vivo with a subcellular resolution that permits the analysis of Ca2+ dynamics down to the level of individual dendritic spines and mitochondria. Here, two available genetically encoded indicators with different Ca2+ affinities are used to demonstrate the use of this protocol for measuring relative Ca2+ levels within the cytoplasm or mitochondrial matrix in a single pair of excitatory interneurons (AVA). Together with the genetic manipulations and longitudinal observations possible in C. elegans, this imaging protocol may be useful for answering questions regarding how Ca2+ handling regulates neuronal function and plasticity.

Subcellular Imaging of Neuronal Calcium Handling In Vivo

The current methods describe a non-ratiometric approach for high-resolution, sub-compartmental calcium imaging in vivo in Caenorhabditis elegans using readily available genetically encoded calcium indicators.

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a ...

Visualizing Proteins in Primary Hippocampal Neurons by Live&#45;Cell Imaging

Using Live&#45;Cell Imaging to Measure the Effects of Pathological Proteins on Axonal Transport in Primary Hippocampal Neurons

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal transport is disrupted in multiple neurodegenerative diseases, and projection neurons are particularly vulnerable because of the need to transport cellular materials over long distances and sustain substantial axonal mass. Pathological modifications of several disease-related proteins negatively affect transport, including tau, amyloid-&#946;, &#945;-synuclein, superoxide dismutase, and huntingtin, providing a potential common mechanism by which pathological proteins exert toxicity in disease. Methods to study these toxic mechanisms are necessary to understand neurodegenerative disorders and identify potential therapeutic interventions.
Here, cultured primary rodent hippocampal neurons are co-transfected with multiple plasmids to study the effects of pathological proteins on fast axonal transport using live-cell confocal imaging of fluorescently-tagged cargo proteins. We begin with the harvest, dissociation, and culturing of primary hippocampal neurons from rodents. Then, we co-transfect the neurons with plasmid DNA constructs to express fluorescent-tagged cargo protein and wild-type or mutant tau (used as an exemplar of pathological proteins). Axons are identified in live cells using an antibody that binds an extracellular&#160;domain of neurofascin, an axon initial segment protein, and an axonal region of interest is imaged to measure fluorescent cargo transport.
Using KymoAnalyzer, a freely available ImageJ macro, we extensively characterize the velocity, pause frequency, and directional cargo density of axonal transport, all of which may be affected by the presence of pathological proteins. Through this method, we identify a phenotype of increased cargo pause frequency associated with the expression of pathological tau protein. Additionally, gene-silencing shRNA constructs can be added to the transfection mix to test the role of other proteins in mediating transport disruption. This protocol is easily adaptable for use with other neurodegenerative disease-related proteins and is a reproducible method to study the mechanisms of how those proteins disrupt axonal transport.

Author Spotlight: Understanding the Impact of Pathological Proteins on Axonal Transport in Neurodegenerative Diseases

Here, we demonstrate how to combine transfection of primary hippocampal rodent neurons with live-cell confocal imaging to analyze pathological protein-induced effects on axonal transport and identify mechanistic pathways mediating these effects.

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

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