The authors thank Dr. James Bamburg and members of his laboratory for collaboration in these studies. This work was funded by NIH grants HD19950, EY07133 and by grants from the Minnesota Medical Foundation.

For rhodamine-actin labeling, neurons are cultured on glass coverslips placed in the bottom of 35 mm plastic dishes, or on coverslips glued into "video" dishes.
1. Preparation of Coverslips or "Video" Dishes
<ol>
<li>Many neuronal types are poorly adhesive to in vitro substrates. Glass coverslips, which are required for this procedure, may not allow sufficient neuron-substrate adhesion unless they are adequately cleaned and prepared. A detailed procedure for acid washing and cleaning glass coverslips is described in the book Culturing Nerve Cells, edited by Banker and Goslin. Alternatively, a nitrocellulose coating for coverslips, which avidly binds substrate molecules, is described in steps 1.7 and 1.8 below.</li>
<li>Glass coverslips (e.g. 18 mm squares or 24 mm circles) are rinsed in dH2O and baked in dry heat (&ge;225&deg;C) as long as possible (minimum of several days up to three months or longer) to remove traces of organic compounds. Go to step 1.6 or for video dishes proceed to 1.3.</li>
<li>To create a thin clear glass substrate for high resolution videomicroscopy, circular holes of appropriate diameter are drilled into the bottoms of 35 or 50 mm plastic Petri or tissue culture dishes using an electric cork borer (or similar instrument). Drilling is done slowly with light pressure to avoid cracking the plastic. Edges of the drilled hole are scraped with scissors to create a smooth surface on both sides. </li>
<li>Glass coverslips are cleaned for tissue culture use, as described in 1.1 or 1.2. Coverslips are glued in place over the holes, using non-toxic silicone aquarium cement (for 18x18mm coverslips, a 12mm diameter bit is used). The cement is cured for 24 hours. </li>
<li>(The following steps are done using sterile conditions) The video dishes are rinsed 3 times with sterile dH2O and allowed to air dry.</li>
<li>Coverslips are coated with substrate molecules for neuronal culture. First, the coverslip surface is coated with (250 &mu;L for 18x18 coverslip) a solution of 100 &mu;g/ mL poly-D-lysine in PBS for 4-8 hrs (or overnight) in a humidified 38&deg; C incubator. Coverslips are then rinsed 3 times with sterile dH2O to remove unbound poly-D-lysine and allowed to air dry. Many neuronal types are cultured on coverslips coated with poly-D-lysine alone, if serum proteins are included in the culture medium. However, results more relevant to in vivo conditions are obtained by using natural substrate molecules. Solutions of these molecules, such as laminin, fibronectin or L1 CAM (1-20 &mu;g/ mL in PBS), can be directly applied to poly-D-lysine coated coverslips for 4-8 hrs or overnight, but in our lab we routinely coat the prepared coverslips with a thin film of nitrocellulose before applying cell adhesion molecules to increase protein-substrate binding.</li>
<li>Make a 1% nitrocellulose solution by dissolving 5 g nitrocellulose in 5 mL of 100% amyl acetate. Apply 10 &mu;L of this solution to the coverslip and gently spread it over the surface. Air dry 10 minutes.</li>
<li>Apply 250 &mu;L of a solution of 25 &mu;g/ mL laminin or 4 &mu;g/ mL L1 CAM in PBS and spread over the nitrocellulose surface. Incubate 4 hours (or overnight) in a humidified 38&deg; C incubator, aspirate substrate and immediately add culture medium, so the substrate does not dry.</li>
</ol>
2. Preparation of Neuronal Cultures
<ol>
<li>Embryonic day 7 chick embryos are removed from eggs, and placed in 100 mm petri dishes containing culture medium with 10% serum for dissection to remove the internal organs from the thorax and abdomen, using a stereomicroscope.</li>
<li>The embryos are then rinsed with medium and moved to a 60 mm dish with liquid medium and further dissected to remove and prepare explants of dorsal root ganglia (DRG) or neural retinal tissues. A detailed dissection description can be found in Methods in Cell Biology, volume 71, Neurons: Methods and Applications for the Cell Biologist, edited by Hollenbeck and Bamburg. DRG explants are 1/2-whole ganglia and neural retinal explants are 1 mm or less in diameter.</li>
<li>After cleaning the explants of extraneous tissues, individual explants are moved with watchmakers forceps to the coverslips, which are already in culture dishes with the culture medium. Under a dissecting microscope, the explant is gently pressed to the center of the coverslip with forceps to prevent movement while transporting to the incubator. Explants are cultured in F12 medium with B27 additives, buffered with 10 mM HEPES to pH 7.4 and placed overnight in a humidified 38&deg; C incubator. Growth factors can be added, as needed, to the culture medium. Neurotrophins are often added to DRG cultures from older embryos, although E7 DRG neurons will extend axons from explants without added neurotrophins. Explants of neural retina extend axons in this culture medium without adding growth factors.</li>
<li>Cultures are incubated for at least 18 hours to allow sufficient axonal outgrowth before beginning the procedure below.</li>
</ol>
3. Preparation of Rhodamine-actin Permeabilization Buffer
<ol>
<li>A stock solution of permeabilization buffer containing 138 mM KCl, 10 mM PIPES, 3 mM EGTA, 4mM MgCl2, and 1% BSA (pH = 6.9) can be kept up to 2 weeks at 4&deg;C11. Just before use, the following components are added for a final concentration of: 0.025% saponin, 0.1 mM ATP, and 100 nM Alexa-fluor 350 phalloidin.</li>
<li>A separate solution is also freshly prepared just before use with these same components plus 0.45&mu;M rhodamine non-muscle actin The solution is alternately vortexed and tritrated for 5 min to dissolve rhodamine-actin clumps, and is vortexed for 1 min during the initial permeabilization step to prevent polymerization in the tube. If despite this, filament assembly in the tube remains a problem, the solution can be centrifuged prior to use.</li>
</ol>
4. Permeabilization of Neuronal Cultures and Incorporation of Rhodamine-actin onto F-actin Barbed Ends
<ol>
<li>A culture dish with DRG or retinal explants is carefully removed from the incubator and the following steps are performed at room temperature.</li>
<li>All changes of solutions must be done carefully. The pipette tip should be placed at the edge of the coverslip, and solutions should be removed and added slowly and with minimal force.</li>
<li>The culture medium is removed by pipette carefully, but steadily, and enough permeabilization buffer to cover cells is added for 1 min (for 18mm x 18mm coverslip, ~60 &mu;L). The culture is not rinsed with any other solutions before adding the permeabilization buffer, and the coverslip is not allowed to dry before adding the permeabilization buffer.</li>
<li>The permeabilization buffer is gently removed by pipette and replaced with the permeabilization buffer containing 0.45 &mu;M rhodamine non-muscle actin for 4 min.</li>
<li>The rhodamine-actin-containing buffer is gently removed by pipette, and the neurons are fixed by adding a solution of 4% paraformaldehyde (laboratory grade, made with phosphate buffer, pH 7.3), 0.05% glutaraldehyde and 10% sucrose. After 5 minutes fixation the coverslips are gently rinsed with PBS, and mounted in Slowfade on 3-inch glass slides. </li>
<li>An alternative approach to changing solutions is to use a flow cell. This can be a commercially available flow cell, or a simple flow cell can be made by placing small pieces of plastic (&le; 1 mm thick) as spacers at the corners of the coverslip, and then mounting an equal-sized coverslip on top of the coverslip. Solutions can be exchanged by placing the pipette at one side of the flow cell and withdrawing solutions with a wick (cotton or a Kimwipe) at the other side. </li>
<li>The incorporation of rhodamine-actin onto F-actin barbed ends and fluorescent-phalloidin labeling of F-actin in growth cones is visualized with epi-fluorescence optics through a 60X oil immersion objective, and images are collected, using a cooled CCD camera. A confocal microscope may provide improved imaging.</li>
<li>For best quantification of rhodamine-actin incorporation all images should be collected in a single session, using the same gain and exposure settings of the digital camera sensitivity for each image. Analysis of labeling should be made from equivalent regions of each image; Metamorph, Image J or similar computerized tools for image analysis can be used.</li>
</ol>
VARIATIONS OF THIS PROCEDURE
5. Variation One: Collect Live Cell Images Prior to Rhodamine-actin Labeling
<ol>
<li>Video dishes are placed onto a warmed microscope stage, and live growth cone images are collected. Gridded coverslips can be used, or etchings and/or pen markings on the coverslip can be made to more easily find specific cells after labeling, or the video dish can be fixed in place on the microscope stage for the duration of the procedure.</li>
<li>Permeabilization, incorporation of rhodamine-actin, and cell fixation are conducted in the video dish, either on or off the microscope stage. After fixing, PBS is added to video dishes for immediate imaging. To preserve the sample for later imaging, slowfade mounting medium is added to the coverslip and a coverslip of equal size is gently placed on top (avoiding bubbles), and the edges are then sealed with clear nail polish.</li>
</ol>
6. Variation Two: Immunochemistry to Co-label Additional Proteins
<ol>
<li>Rhodamine-actin incorporation and fixation can be conducted on neurons cultured on coverslips or in video dishes, as described above (4.2-4.4).</li>
<li>After fixation, and rinsing in PBS, cells are treated 15 min with 0.1 M glycine in PBS and then extracted with 0.1% Triton X-100 (TX-100) in PBS with 2% goat serum and 1% BSA for 1 h. Coverslips are incubated with the primary antibodies diluted in PBS containing 1% BSA for 1 h. The coverslips are then rinsed and incubated in 0.1%TX-100 in PBS with 2% goat serum and 1% BSA for 1 h, before applying fluorescent secondary antibodies at 1:1000 dilution in PBS with 1% BSA for 1 h. After rinsing, the coverslips are again incubated in 0.1% TX-100 in PBS with 2% goat serum and 1% BSA for 30 min, rinsed, and mounted in anti-fading medium. The localization of some proteins is disrupted by the initial permeabilization step (4.3). To retain these proteins for localization with antibodies, 0.05% paraformaldehyde and 0.05% glutaraldehyde can be added to the permeabilization buffer for 1 min (i.e. before adding the rhodamine-actin) without affecting the barbed end labeling by rhodamine-actin. </li>
<li>Images of triple-labeled growth cones are collected by fluorescence or confocal microscopy.</li>
</ol>
7. Variation Three: Assessment of the Effects of Guidance Cues on F-actin Free Barbed Ends
<ol>
<li>A growth factor or guidance cue is added to culture dishes in the incubator at concentrations of several ng/ mL for a few minutes (or an appropriate treatment time) before removing the dish from the incubator and beginning the permeabilization procedure. This allows assessment of the short-term effects of growth factors or guidance cues on F-actin free barbed ends.</li>
<li>A video dish with neuronal cultures can be placed on a warmed microscope stage, and growth factors or guidance cues can be released from a micropipette in a gradient near a growth cone before beginning the permeabilization and rhodamine-actin incorporation. For example, NGF can be released for DRG neurons or netrin for neural retinal ganglion cells. The pipette can be introduced for as short as 1-2 minutes before beginning the permeabilization procedure. This allows assessment of the local effects of a guidance cue gradient on the formation of F-actin free barbed ends in growth cone leading margins (see reference citation 10 for images).</li>
<li>These variations can be combined with antibody-mediated labeling of other neuronal components, as described in 6.2 above.</li>
</ol>
8. Variation Four: F-actin Free Barbed End Labeling on Transfected Neurons
<ol>
<li>The use of constitutively active or dominant-negative constructs or protein knockdown using RNAi (with a fluorescent identifying marker) can be used to assess a protein's involvement in regulating F-actin free barbed ends. Depending on the protein, its localization within the growth cone, and if it is fused to a fluorescent tag, the fluorescent expressed protein in transfected cells may be lost upon permeabilization. If this is the case, video dishes can be used to identify a transfected cell microscopically before permeabilization, and the permeabilization can be performed on the microscope stage, after marking the positions of transfected cells. Alternatively, a GFP-actin construct can be co-transfected with a GFP-construct of interest. In this case, GFP-actin will be incorporated into F-actin, and will not be lost by permeabilization, enabling the identification of transfected cells after permeabilization. </li>
<li>This variation can be combined with the addition of guidance cues, as described in 7.1-7.3 above.</li>
</ol>
9. Representative Results
<img src="/files/ftp_upload/2409/2409fig1.jpg" alt="Figure 1" /> Figure 1. Global addition of NGF increases total F-actin and F-actin barbed ends at the growth cone leading margin. When a DRG growth cone is stimulated with nerve growth factor (NGF) for 5 minutes, actin polymerization is stimulated at the leading margin, and a bright band of rhodamine-actin labeling is seen around the growth cone periphery. Figure 1 compares rhodamine-actin incorporation into an unstimulated DRG growth cone and an NGF-stimulated DRG growth cone. The green fluorescence in the merged images is phalloidin labeling for F-actin in the growth cone and the red is rhodamine-actin labeling. A good control for the barbed end labeling is to add 10-6 M or higher cytochalasin B or D in the permeabilization buffer in steps 4.2 and 4.3. Cytochalasins cap F-actin barbed ends and inhibit rhodamine-actin binding to barbed ends. This should severely reduce or eliminate incorporation of rhodamine-actin into the cell. Scale bars, 10 &mu;m.
<img src="/files/ftp_upload/2409/2409fig2.jpg" alt="Figure 2" /> Figure 2. An NGF gradient locally increases F-actin barbed ends. A micropipette that releases NGF is brought to one side of a DRG growth cone for 2 minutes, followed by labeling of F-actin free barbed ends. The images below show that exposure to a gradient of NGF locally stimulates an increase in F-actin free barbed ends in the growth cone region closer to the pipette. The merged image shows phalloidin in green and rhodamine-actin in red. Scale bar, 10 &mu;m.
<img src="/files/ftp_upload/2409/2409fig3.jpg" alt="Figure 3" /> Figure 3. Activated ERM proteins accumulate at the growth cone leading margin. Immunocytochemical labeling of phospho-Ezrin/Radixin/Moesin (pERM) with F-actin free barbed ends. Gluteraldehyde (0.05%) and paraformaldehyde (0.05%) were added to the initial permeabilization buffer for one min to preserve ERM localization. Rhodamine-actin, red; pERM, green; phalloidin, blue. Scale bar, 10 &mu;m.
<img src="/files/ftp_upload/2409/2409fig4.jpg" alt="Figure 4" /> Figure 4. Stimulation of actin polymerization requires active ERM proteins. Dissociated DRGs were co-transfected with GFP-actin and a GFP control plasmid or a dominant-negative Ezrin/Radixin/Moesin (DN ERM) construct. The use of GFP-actin allows identification of transfected cells after permeabilization. Here, NGF was added 5 min prior to labeling of F-actin barbed ends. Note the growth cone transfected with DN ERM has reduced barbed end levels at the growth cone leading margin (lower middle panel), which contrasts with a nearby untransfected growth cone (lower panels), or with the GFP control (upper middle panel). Rhodamine-actin, red; GFP-actin, green. Scale bars, 10 &mu;m.

<ol>
	<li>Lowery, L. A., Van Vactor, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+trip+of+the+tip:+Understanding+the+growth+cone+machinery.">The trip of the tip: Understanding the growth cone machinery.</a> Nat Rev Mol Cell Biol. 10, 332-343 (2009).</li><li>Letourneau, P., Squire, L. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axonal+Pathfinding:+Extracellular+Matrix+Role.">Axonal Pathfinding: Extracellular Matrix Role.</a> Encyclopedia of Neuroscience. , 1139-1145 (2008).</li><li>Kalil, K., Dent, E. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Touch+and+go:+Guidance+cues+signal+to+the+growth+cone+cytoskeleton.">Touch and go: Guidance cues signal to the growth cone cytoskeleton.</a> Curr Opin Neurobio. 15, 521-526 (2005).</li><li>Dent, E. W., Gertler, F. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoskeletal+dynamics+and+transport+in+growth+cone+motility+and+axon+guidance.">Cytoskeletal dynamics and transport in growth cone motility and axon guidance.</a> Neuron. 40, 209-227 (2003).</li><li>Pak, C. W., Flynn, K. C., Bamburg, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin-binding+proteins+take+the+reins+in+growth+cones.">Actin-binding proteins take the reins in growth cones.</a> Nat Rev Neurosci. 9, 136-147 (2008).</li><li>Guan, K. L., Rao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signaling+mechanisms+mediating+neuronal+responses+to+guidance+cues.">Signaling mechanisms mediating neuronal responses to guidance cues.</a> Nat Rev Neurosci. 4, 941-956 (2003).</li><li>Gallo, G., Letourneau, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+growth+cone+actin+filaments+by+guidance+cues.">Regulation of growth cone actin filaments by guidance cues.</a> J. Neurobiology. 58, 92-102 (2004).</li><li>Fass, J., Gehler, S., Sarmiere, P., Letourneau, P., Bamburg, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulating+filopodial+dynamics+through+actin-depolymerizing+factor/cofilin.">Regulating filopodial dynamics through actin-depolymerizing factor/cofilin.</a> Anat Sci Inter. 79, 173-183 (2004).</li><li>Bernstein, B. W., Bamburg, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ADF/cofilin:+A+functional+node+in+cell+biology.">ADF/cofilin: A functional node in cell biology.</a> Trends Cell Biol. 20, 187-195 (2010).</li><li>Marsick, B. M., Flynn, K. C., Santiago, M., Bamburg, J. R., Letourneau, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+ADF/cofilin+mediates+attractive+growth+cone+turning+toward+nerve+growth+factor+and+netrin-1.">Activation of ADF/cofilin mediates attractive growth cone turning toward nerve growth factor and netrin-1.</a> Dev Neurobiol. 70, 565-588 (2010).</li><li>Symons, M. H., Mitchison, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+actin+polymerization+in+live+and+permeabilized+fibroblasts.">Control of actin polymerization in live and permeabilized fibroblasts.</a> J Cell Biol. 114, 503-513 (1991).</li><li>Chan, A. Y., Raft, S., Bailly, M., Wyckoff, J. B., Segall, J. E., Condeelis, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EGF+stimulates+an+increase+in+actin+nucleation+and+filament+number+at+the+leading+edge+of+the+lamellipod+in+mammary+adenocarcinoma+cells.">EGF stimulates an increase in actin nucleation and filament number at the leading edge of the lamellipod in mammary adenocarcinoma cells.</a> J Cell Sci. 111, 199-211 (1998).</li></ol>

No conflicts of interest declared.

The methods presented here allow temporal and spatial resolution of cellular components that are involved in the dynamic remodeling of the actin cytoskeleton at the leading margin of migrating growth cones. The action of attractant molecules, like NGF or netrin, to rapidly stimulate actin filament polymerization is revealed as a local increase in actin barbed ends, created by actin filament severing by activated ADF/cofilin10, as shown in Figures 1 and 2. The method permits localization of other proteins invol...

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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
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<tr>
<td>18x18 mm coverslip</td>
<td>&nbsp;</td>
<td>Gold Seal</td>
<td>3305</td>
<td>&nbsp;</td>
<tr>
<td>35mm Petri dishes</td>
<td>&nbsp;</td>
<td>Falcon</td>
<td>351008</td>
<td>&nbsp;</td>
<tr>
<td>Aquarium cement</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>DAP 100% silicone aquarium sealant</td>
<td>Any hardware store</td>
</tr>
<tr>
<td>Poly-D-lysine (mw &gt; 300,000)</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P1024</td>
<td>&nbsp;</td>
<tr>
<td>Natural mouse laminin</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>23017-015</td>
<td>&nbsp;</td>
<tr>
<td>L1 CAM</td>
<td>&nbsp;</td>
<td>R &amp; D systems</td>
<td>777-NC</td>
<td>&nbsp;</td>
<tr>
<td>Alexa-fluor 350 phalloidin</td>
<td>&nbsp;</td>
<td>Invitrogen (Molecular Probes)</td>
<td>A22281</td>
<td>&nbsp;</td>
<tr>
<td>Rhodamine non-muscle actin</td>
<td>&nbsp;</td>
<td>Cytoskeleton, Inc.</td>
<td>APHR-A</td>
<td>&nbsp;</td>
<tr>
<td>F12 Culture medium</td>
<td>&nbsp;</td>
<td>Invitrogen (Gibco)</td>
<td>21700-075</td>
<td>&nbsp;</td>
<tr>
<td>B27</td>
<td>&nbsp;</td>
<td>Invitrogen (Gibco)</td>
<td>17504-044</td>
<td>&nbsp;</td>
<tr>
<td>Slowfade </td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>536937</td>
<td>&nbsp;</td>		</tbody>
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labeling f-actin barbed ends with rhodamine-actin in permeabilized neuronal growth cones

The motile tips of growing axons are called growth cones. Growth cones lead navigating axons through developing tissues by interacting with locally expressed molecular guidance cues that bind growth cone receptors and regulate the dynamics and organization of the growth cone cytoskeleton3-6. The main target of these navigational signals is the actin filament meshwork that fills the growth cone periphery and that drives growth cone motility through continual actin polymerization and dynamic remodeling7. Positive or attractive guidance cues induce growth cone turning by stimulating actin filament (F-actin) polymerization in the region of the growth cone periphery that is nearer the source of the attractant cue. This actin polymerization drives local growth cone protrusion, adhesion of the leading margin and axonal elongation toward the attractant.
Actin filament polymerization depends on the availability of sufficient actin monomer and on polymerization nuclei or actin filament barbed ends for the addition of monomer. Actin monomer is abundantly available in chick retinal and dorsal root ganglion (DRG) growth cones. Consequently, polymerization increases rapidly when free F-actin barbed ends become available for monomer addition. This occurs in chick DRG and retinal growth cones via the local activation of the F-actin severing protein actin depolymerizing factor (ADF/cofilin) in the growth cone region closer to an attractant8-10. This heightened ADF/cofilin activity severs actin filaments to create new F-actin barbed ends for polymerization. The following method demonstrates this mechanism. Total content of F-actin is visualized by staining with fluorescent phalloidin. F-actin barbed ends are visualized by the incorporation of rhodamine-actin within growth cones that are permeabilized with the procedure described in the following, which is adapted from previous studies of other motile cells11, 12. When rhodamine-actin is added at a concentration above the critical concentration for actin monomer addition to barbed ends, rhodamine-actin assembles onto free barbed ends. If the attractive cue is presented in a gradient, such as being released from a micropipette positioned to one side of a growth cone, the incorporation of rhodamine-actin onto F-actin barbed ends will be greater in the growth cone side toward the micropipette10.
Growth cones are small and delicate cell structures. The procedures of permeabilization, rhodamine-actin incorporation, fixation and fluorescence visualization are all carefully done and can be conducted on the stage of an inverted microscope. These methods can be applied to studying local actin polymerization in migrating neurons, other primary tissue cells or cell lines.

Watch this Scientific Journal Video about Labeling F-actin Barbed Ends with Rhodamine-actin in Permeabilized Neuronal Growth Cones at JoVE.com

Labeling F-actin Barbed Ends with Rhodamine-actin in Permeabilized Neuronal Growth Cones

A method to visualize and quantify F-actin barbed ends in neuronal growth cones is described. After culturing neurons on glass coverslips, cells are permeabilized with a saponin-containing solution. Then, a short incubation with the saponin buffer containing rhodamine-actin incorporates fluorescent actin onto free actin barbed ends.

The motile tips of growing axons are called growth cones. Growth cones lead navigating axons through developing tissues by interacting with locally ...

labeling-f-actin-barbed-ends-with-rhodamine-actin-permeabilized

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Neuroscience

14.7K vistas.  University of Minnesota. Un método para visualizar y cuantificar la F-actina fines de púas en los conos de crecimiento neuronal se describe. Después de las neuronas de cultivo en cubreobjetos de vidrio, las células se permeabilized con una solución de saponina que contiene. A continuación, un corto de incubación con el tampón de saponina que contiene rodamina-actina incorpora actina fluorescente en los extremos de actina de púas libre.

Video: Labeling F-actin Barbed Ends with Rhodamine-actin in Permeabilized Neuronal Growth Cones

DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; Gan et al., 2000). Here we provide a detailed description of each step required together with exemplary images of good and bad outcomes that will help when setting up the technique. In our experience, a few steps proved critical for the successful application of DiOLISTICS. These considerations include the quality of the DiI-coated bullets, the extent of fixative exposure, and the concentration of detergent used in the incubation solutions. Tips and solutions for common problems are provided. This is a versatile labeling technique that can be applied to multiple animal species at a wide range of ages. Unlike other fluorescent labeling techniques that are limited to preparations from young animals or restricted to mice because they rely on the expression of a fluorescent transgene, DiOLISTIC labeling can be applied to animals of all ages, species and genotypes and it can be used in combination with immunostaining to identify a specific subpopulation of cells. Here we demonstrate the use of DiOLISTICS to label neurons in brain slices from adult mice and adult non-human primates with the purpose of quantifying dendrite branching and dendritic spine morphology.

We demonstrate the use of the gene gun to introduce fluorescent dyes, such as DiI, into neurons in brain slices from rodents and non-human primates of different ages. In this particular case, we use adult mice (3-6 months old) and adult cynomologus monkeys (9-15 years old). This technique, originally described by the laboratory of Dr. Lichtman (Gan et al., 2000), is well suited for the study of dendritic branching and dendritic spine morphology and can be combined with traditional immunostaining, if detergents are kept at a low concentration.

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; ...

Automated Sholl Analysis of Digitized Neuronal Morphology at Multiple Scales

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3. Specifically, it affects the ability of neurons to receive inputs from other cells2 and contributes to the propagation of action potentials4,5. The morphology of the neurites also affects how information is processed. The diversity of dendrite morphologies facilitate local and long range signaling and allow individual neurons or groups of neurons to carry out specialized functions within the neuronal network6,7. Alterations in dendrite morphology, including fragmentation of dendrites and changes in branching patterns, have been observed in a number of disease states, including Alzheimer's disease8, schizophrenia9,10, and mental retardation11. The ability to both understand the factors that shape dendrite morphologies and to identify changes in dendrite morphologies is essential in the understanding of nervous system function and dysfunction.
Neurite morphology is often analyzed by Sholl analysis and by counting the number of neurites and the number of branch tips. This analysis is generally applied to dendrites, but it can also be applied to axons. Performing this analysis by hand is both time consuming and inevitably introduces variability due to experimenter bias and inconsistency. The Bonfire program is a semi-automated approach to the analysis of dendrite and axon morphology that builds upon available open-source morphological analysis tools. Our program enables the detection of local changes in dendrite and axon branching behaviors by performing Sholl analysis on subregions of the neuritic arbor. For example, Sholl analysis is performed on both the neuron as a whole as well as on each subset of processes (primary, secondary, terminal, root, etc.) Dendrite and axon patterning is influenced by a number of intracellular and extracellular factors, many acting locally. Thus, the resulting arbor morphology is a result of specific processes acting on specific neurites, making it necessary to perform morphological analysis on a smaller scale in order to observe these local variations12.
The Bonfire program requires the use of two open-source analysis tools, the NeuronJ plugin to ImageJ and NeuronStudio. Neurons are traced in ImageJ, and NeuronStudio is used to define the connectivity between neurites. Bonfire contains a number of custom scripts written in MATLAB (MathWorks) that are used to convert the data into the appropriate format for further analysis, check for user errors, and ultimately perform Sholl analysis. Finally, data are exported into Excel for statistical analysis. A flow chart of the Bonfire program is shown in Figure 1.

We have developed a computer program to analyze neuronal morphology. In combination with two existing open source analysis tools, our program performs Sholl analysis and determines the number of neurites, branch points, and neurite tips. The analyses are performed so that local changes in neurite morphology can be observed.

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3.  Specifically, it affects the ability of neurons to ...

Recording Large-scale Neuronal Ensembles with Silicon Probes in the Anesthetized Rat

Large scale electrophysiological recordings from neuronal ensembles offer the opportunity to investigate how the brain orchestrates the wide variety of behaviors from the spiking activity of its neurons. One of the most effective methods to monitor spiking activity from a large number of neurons in multiple local neuronal circuits simultaneously is by using silicon electrode arrays1-3.
Action potentials produce large transmembrane voltage changes in the vicinity of cell somata. These output signals can be measured by placing a conductor in close proximity of a neuron. If there are many active (spiking) neurons in the vicinity of the tip, the electrode records combined signal from all of them, where contribution of a single neuron is weighted by its 'electrical distance'. Silicon probes are ideal recording electrodes to monitor multiple neurons because of a large number of recording sites (+64) and a small volume. Furthermore, multiple sites can be arranged over a distance of millimeters, thus allowing for the simultaneous recordings of neuronal activity in the various cortical layers or in multiple cortical columns (Fig. 1). Importantly, the geometrically precise distribution of the recording sites also allows for the determination of the spatial relationship of the isolated single neurons4. Here, we describe an acute, large-scale neuronal recording from the left and right forelimb somatosensory cortex simultaneously in an anesthetized rat with silicon probes (Fig. 2).

Extracellular recordings of neuronal activity using silicon probes in the anesthetized rat will be described. This technique allows information to be obtained across multiple brain areas from more than 100 neurons simultaneously. It provides information with single cell resolution about neuronal ensembles dynamics in multiple local circuits.

Large scale electrophysiological recordings from neuronal ensembles offer the opportunity to investigate how the brain orchestrates the wide variety of ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

Preparation of Neuronal Co-cultures with Single Cell Precision

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms can be used to investigate a variety of questions, spanning developmental and functional neurobiology to infection and disease propagation. However, conventional systems require significant cellular inputs (many thousands per compartment), inadequate for studying low abundance cells, such as primary dopaminergic substantia nigra, spiral ganglia, and Drosophilia melanogaster neurons, and impractical for high throughput experimentation. The dense cultures are also highly locally entangled, with few outgrowths (&#60;10%) interconnecting the two cultures. In this paper straightforward microfluidic and patterning protocols are described which address these challenges: (i) a microfluidic single neuron arraying method, and (ii) a water masking method for plasma patterning biomaterial coatings to register neurons and promote outgrowth between compartments. Minimalistic neuronal co-cultures were prepared with high-level (&#62;85%) intercompartment connectivity and can be used for high throughput neurobiology experiments with single cell precision.

Protocols for single neuron microfluidic arraying and water masking for the in-chip plasma patterning of biomaterial coatings are described. Highly interconnected co-cultures can be prepared using minimal cell inputs.

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...