The authors thank Teele Palumaa for assistance with the figures. Funded by Project Grant 1008046 from the National Health and Medical Research Council, Australia.

1. Dissociated Hippocampal Neuron Culture
<ol>
	<li>Prepare coverslips (borosilicate glass):
	<ol>
		<li>Wash in 100% ethanol.</li>
		<li>Air dry under UV irradiation.</li>
		<li>Coat with Poly-D-lysine (0.5 mg/ml in 0.15 M borate buffer, overnight at 4 &#176;C).</li>
		<li>On the next day (the day of culture) wash 3x in PBS then coat with laminin (2.5 &#956;g/ml, natural mouse laminin) + 5% v/v heat-inactivated fetal calf serum (FCS) diluted in PBS for 2 hr at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Dissect embryonic day 18 (E18) rat hippocampi and collect into PBS containing calcium and magnesium, chilled on ice. NOTE: all experimental protocols involving animals were approved by the Animal Ethics Committee of the University of Melbourne.</li>
	<li>Prepare the papain and DNase I solutions from the Papain Dissociation Kit according to the manufacturer&#8217;s instructions.
	<ol>
		<li>Add 50 &#956;l DNase I solution to 1 ml papain solution.</li>
		<li>NOTE: Once first prepared, the papain solution and the DNase I solutions may be separately dispensed into aliquots (0.5 ml and 25 &#956;l aliquots for papain and DNase I, respectively) and stored at -20 &#176;C until needed.</li>
	</ol>
	</li>
	<li>Remove as much PBS as possible and incubate hippocampi (from one litter of embryos) with 1 ml papain/DNase I solution at 37 &#176;C for 15-20 min. Gently flick the tube to mix the contents twice during the incubation period.</li>
	<li>Triturate the hippocampal tissue gently (avoiding the generation of bubbles) with a flame-polished siliconized Pasteur pipette 10-15x until cell dispersion is obtained and few, if any, chunks of undissociated tissue remain.</li>
	<li>Carefully layer the dissociated cell suspension over a 3 ml cushion of 4% w/v bovine serum albumin (BSA) in Hanks Balanced Salt Solution plus additives (HBSS+; see Section 1.6.1).
	<ol>
		<li>Prepare this step gradient solution by dissolving the BSA at room temperature without stirring in HBSS containing 2 mM CaCl2, 1 mM MgSO4, 4 mM NaHCO3 and 40 mM glucose (HBSS+) then sterile filter and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Centrifuge, 100 x g, 7 min in a bench top centrifuge with a swing-out rotor.</li>
	<li>Remove the supernatant being careful not to aspirate the cell pellet.</li>
	<li>Resuspend the pelleted cells gently with a flame-polished (siliconized) Pasteur pipette (or a 1 ml blue pipette tip) in 1 ml complete Neurobasal medium (with 2% B27, 0.5 mM L-glutamine supplemented with 1% FCS).</li>
	<li>Count two 10 &#956;l aliquots using a hemocytometer (N.B. only count phase-bright cells as &#8220;live&#8221; cells).</li>
	<li>Plate primary neurons on preprepared coated glass coverslips (0.75-1 x 105/18 mm coverslip in 12-well plate). Immediately prior to plating, aspirate excess laminin/serum solution from coverslips and replace with primary neuron culture medium (see below). N.B. The required number of coverslips needs to be determined in advance as coverslip preparation is begun the day prior to culture (see step 1.1).</li>
	<li>Culture rat primary E18 hippocampal neurons for up to 21 days in vitro (DIV) in Neurobasal medium, 2% B27, 0.5 mM L-glutamine supplemented with 1% FCS (heat inactivated). Perform a half medium change at 7 DIV and weekly thereafter. The anti-mitotic fluorodeoxyuridine/uridine is added at 7 DIV; 1/1,000 dilution of 10 mM stock of each nucleoside) to prevent glial overgrowth.</li>
</ol>
2. Antibody Incubation with Live Neurons
<ol>
	<li>During the first week in culture, neurons are developing dendritic arbors and by week 2-3, neurons have matured sufficiently to be undergoing synaptogenesis6,7.
	<ol>
		<li>At selected experimental time point/s, apply the primary antibody to triplicate wells containing primary embryonic neurons on coverslips. Add an aliquot of the antibody directly into the culture medium of the primary neurons to the desired final dilution Note: In our example, a rabbit polyclonal antiserum raised against a recombinant secreted form of the protein was diluted 1/500 by adding 2 &#956;l to 1 ml of culture medium in well; a centrifugation for 10 min, 13,000 x g, RT can be incorporated prior to taking an aliquot of antibody as this will remove any particulate matter and can help reduce background.</li>
		<li>For the preimmune serum controls, add an equivalent amount of preimmune serum (to the same final dilution). If no preimmune serum is available, add the equivalent volume of Neurobasal medium or PBS (no primary antibody control). Alternatively, if a suitable primary antibody is available that recognizes intracellular region/s of the protein, this antibody can be added to a control well in order to test specificity of the surface staining.</li>
		<li>Return the cells to the culture incubator for 1-4 hr (this incubation period may be determined empirically). In our experiments, the antibody was present for the entire period although the experimental design could incorporate a pulse of antibody followed by a further incubation period after wash-out (a pulse-chase experiment) to assess the time course of internalization. 
		OPTIONAL STEP: This incubation can be carried out at room temperature or even on ice to slow the rate of basal protein internalization, if necessary. If performed in a non-CO2 controlled environment, the medium should be changed to one that is not bicarbonate buffered before addition of the antibody/antiserum. CAUTION: Mature neuron cultures (&#62;14 days, and particularly cultured mouse embryonic neurons) do not tolerate full medium changes well.</li>
	</ol>
	</li>
	<li>We have used incubation times for this primary antibody internalization step of 1 hr, 2 hr, and 4 hr with good results although the optimal time will depend on abundance of the protein of interest as well as the dynamics of protein trafficking in the cells under study. 
	NOTE: The dual-color images shown in Representative Results were obtained with an incubation of 1 hr at 37 &#176;C in a tissue culture incubator.</li>
	<li>After incubation for the desired period, aspirate off the antibody-containing medium. Wash the wells gently but rapidly once with PBS at room temperature.</li>
</ol>
3. Secondary Antibody Application to Fixed, Unpermeabilized Cells
<ol>
	<li>Fix neurons with 4% paraformaldehyde in phosphate buffer pH 7.2, 5 min at room temperature Note: for best results, use freshly prepared fixative; alternatively use fix that has been stored at -20 &#176;C and completely thawed so that no precipitated paraformaldehyde is evident. CAUTION: Paraformaldehyde fixative should be prepared and handled in a fume hood.</li>
	<li>Remove fix from wells (transfer to liquid waste container in fume hood) and rinse 3x with PBS.
	<ol>
		<li>OPTIONAL STEP: If desired, to save on antibody reagents, the coverslips can be carefully removed from the 12-well plate with forceps and placed cell-side-up, onto a sheet of Parafilm laid on the base of a disposable culture plate.</li>
		<li>Solutions can then be pipetted gently onto the coverslips so that they are completely covered without the solution flooding over the edge of the coverslip onto the Parafilm.</li>
		<li>This technique is suitable for short-term incubations (1-2 hr) however longer incubations (e.g. overnight) should be carried out in a humidified chamber. Alternatively, the coverslips can be placed back into the wells of the 12-well plate for the overnight incubation and sufficient volume should be added to ensure that the coverslips do not dry out.</li>
	</ol>
	</li>
	<li>Block for 30 min&#160;at room temperature with 5% BSA in PBS (Note:&#160;Do not add detergent to the blocking solution at this stage as it is important that the cells are not permeabilized).</li>
	<li>In order to label surface protein before proceeding with detection of internalized protein, apply the first fluorescently labeled 2&#176; antibody of choice. 
	Note: In the example described here, the primary antiserum was raised in rabbit (in-house antibody) to a recombinant secreted isoform4. Thus, for the first secondary antibody to detect the extracellular region of the transmembrane isoform on the surface of unpermeabilized neurons, we used donkey anti-rabbit Dylight 649 (1/200 diluted in PBS containing 5% BSA). It is recommended to centrifuge diluted secondary antibody solutions (10 min, 13,000 x g, RT) prior to use.</li>
	<li>Incubate the coverslips for 2 hr at room temperature.</li>
	<li>Wash coverslips (in wells), 2x 5 min with PBS.</li>
</ol>
4. Blocking with Excess Unlabeled 2&#176; Antibody
<ol>
	<li>Block the unpermeabilized neurons with a high concentration (&#62;0.1 mg/ml) of unlabeled 2&#176; antibody.
	<ol>
		<li>The unlabeled 2&#176; antibody should be raised against the species in which the primary antibody was raised (in this case rabbit) by incubation overnight at room temperature.</li>
		<li>For this protocol, AffiniPure Fab fragment Goat anti-Rabbit IgG (H+L) was used at a concentration of 0.13 mg/ml. 
		NOTE: The overnight incubation was found to be crucial as a shorter incubation period (2 hr) was insufficient for complete blocking of primary antibody that was not fully bound by the labeled secondary antibody.</li>
	</ol>
	</li>
	<li>Wash coverslips (in wells), 2x 5 min with PBS.</li>
	<li>After this blocking step, post-fix the cells with 4% paraformaldehyde in phosphate buffer pH 7.2, 5 min at room temperature. Rinse with PBS (2x) after removal of fixative (transfer fixative to liquid waste container in fume hood).</li>
</ol>
5. Permeabilization and Application of the Second Fluorescently Conjugated 2&#176; Antibody
<ol>
	<li>Permeabilize and block the cells with 5% BSA in PBS containing 0.1% Triton-X-100 at room temperature for 30 min.</li>
	<li>Remove the blocking solution (taking care that the coverslips do not dry out). Add the second fluorescently-conjugated 2&#176; antibody tagged with a different fluorophore.
	<ol>
		<li>NOTE: It must be possible to distinguish this fluorophore tag from the one previously used, depending on the available excitation/emission filters on the confocal microscope (see below).</li>
		<li>For the example presented in this protocol, an Alexa Fluor 488-conjugated donkey anti-rabbit 2&#176; antibody was used (1/200 in PBS, 5% BSA and 0.1% Triton-X-100). Incubate 2 hr at room temperature then remove the 2&#176; antibody solution.</li>
	</ol>
	</li>
	<li>Wash coverslips 3x 5 min with PBS and, finally, wash briefly with deionized water.</li>
</ol>
6. Mounting and Imaging
<ol>
	<li>Mount coverslips on glass slides with an aqueous mounting medium containing antifade (e.g. VECTASHIELD) and allow to dry. Store in the dark at 4 &#176;C for optimal preservation of fluorescent signal intensity.</li>
	<li>Image immunostained cells on a confocal microscope.
	<ol>
		<li>Appropriate excitation and emission filters for detection of the two fluorophore signals must be available.</li>
		<li>If different experimental conditions are to be compared (for example, internalization rates under depolarized8,9 or control conditions), ensure that all replicate coverslips from the various conditions are imaged with the same image acquisition parameters. The integrated density of standard regions of interest or the puncta attributes (e.g. number, size) from the resulting images can then be measured using standard image analysis software (e.g. Fiji/ImageJ, Metamorph).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Lewis, T. L., Mao, T., Arnold, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+role+for+myosin+VI+in+the+localization+of+axonal+proteins.">A role for myosin VI in the localization of axonal proteins.</a> PLoS Biol. 9, (2010).</li><li>von Zastrow, M., Williams, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulating+neuromodulation+by+receptor+membrane+traffic+in+the+endocytic+pathway.">Modulating neuromodulation by receptor membrane traffic in the endocytic pathway.</a> Neuron. 76, 22-32 (2012).</li><li>Gunnersen, J. M., Augustine, C., Spirkoska, V., Kim, M., Brown, M., Tan, S. -. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+analysis+of+gene+expression+patterns+in+developing+mouse+neocortex+using+serial+analysis+of+gene+expression.">Global analysis of gene expression patterns in developing mouse neocortex using serial analysis of gene expression.</a> Mol. Cell. Neurosci. 19, 560-573 (2002).</li><li>Gunnersen, J. M., Kim, M. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sez-6+proteins+affect+dendritic+arborization+patterns+and+excitability+of+cortical+pyramidal+neurons.">Sez-6 proteins affect dendritic arborization patterns and excitability of cortical pyramidal neurons.</a> Neuron. 56, 621-639 (2007).</li><li>Kennedy, M. J., Davison, I. G., Robinson, C. G., Ehlers, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Syntaxin-4+Defines+a+Domain+for+Activity-Dependent+Exocytosis+in+Dendritic+Spines.">Syntaxin-4 Defines a Domain for Activity-Dependent Exocytosis in Dendritic Spines.</a> Cell. 141, 524-535 (2010).</li><li>Cullen, D. K., Gilroy, M. E., Irons, H. R., Laplaca, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synapse-to-neuron+ratio+is+inversely+related+to+neuronal+density+in+mature+neuronal+cultures.">Synapse-to-neuron ratio is inversely related to neuronal density in mature neuronal cultures.</a> Brain Res. 1359, 44-55 (2010).</li><li>Grabrucker, A., Vaida, B., Bockmann, J., Boeckers, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptogenesis+of+hippocampal+neurons+in+primary+cell+culture.">Synaptogenesis of hippocampal neurons in primary cell culture.</a> Cell Tissue Res. , 338-341 (2009).</li><li>Vaillant, A. R., Zanassi, P., Walsh, G. S., Aumont, A., Alonso, A., Miller, F. D. <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+underlying+reversible,+activity-dependent+dendrite+formation.">Signaling mechanisms underlying reversible, activity-dependent dendrite formation.</a> Neuron. 34, 985-998 (2002).</li><li>Park, M., Penick, E. C., Edwards, J. G., Kauer, J. A., Ehlers, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recycling+endosomes+supply+AMPA+receptors+for+LTP.">Recycling endosomes supply AMPA receptors for LTP.</a> Science. 305, 1972-1975 (2004).</li><li>Kennedy, M. J. &. a. m. p. ;., Ehlers, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organelles+and+trafficking+machinery+for+postsynaptic+plasticity.">Organelles and trafficking machinery for postsynaptic plasticity.</a> Annu. Rev. Neurosci. 29, 325-362 (2006).</li><li>Lasiecka, Z. M., Yap, C. C., Caplan, S., Winckler, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+early+endosomes+require+EHD1+for+L1/NgCAM+trafficking.">Neuronal early endosomes require EHD1 for L1/NgCAM trafficking.</a> J. Neurosci. 30, 16485-16497 (2010).</li><li>Arancibia-C&#225;rcamo, I. L., Fairfax, B. P., Moss, S. J., Kittler, J. T., Kittler, J. T., Moss, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studying+the+localization,+surface+stability+and+endocytosis+of+neurotransmitter+receptors+by+antibody+labeling+and+biotinylation+approaches.">Studying the localization, surface stability and endocytosis of neurotransmitter receptors by antibody labeling and biotinylation approaches.</a> The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology. , (2006).</li><li>Petrini, E. M., Lu, J., Cognet, L., Lounis, B., Ehlers, M. D., Choquet, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endocytic+trafficking+and+recycling+maintain+a+pool+of+mobile+surface+AMPA+receptors+required+for+synaptic+potentiation).">Endocytic trafficking and recycling maintain a pool of mobile surface AMPA receptors required for synaptic potentiation).</a> Neuron. 63, 92-105 (2009).</li><li>Reider, A., Wendland, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endocytic+adaptors+&#8211;+social+networking+at+the+plasma+membrane.">Endocytic adaptors &#8211; social networking at the plasma membrane.</a> J. Cell Sci. 124, 1613-1622 (2011).</li></ol>

The authors declare that they have no competing financial interests.

The technique described here is complementary to that of cell-surface biotinylation (reviewed by Arancibia-C&#224;rcamo et al.)12 and it is the method of choice for preserving information about the subcellular localization of the internalized protein, provided a suitable primary antibody to an extracellular epitope is available. In addition, quantitation of protein trafficking/internalization over time can be performed (by fixing coverslips at different times throughout the live cell incubation with p...

In establishing the function of newly identified proteins, investigation of the subcellular localization and trafficking of the protein in question can provide important clues about the likely role/s of the protein1,2. Bioinformatic analysis of the transcriptome of the developing neocortex3 provided us with a list of genes exhibiting altered expression during mouse brain corticogenesis. We then adopted a gene knockout approach to ascertain that the protein encoded by one of these genes, sez6, has a key role in neuron development. We observed that the Seizure-related gene 6, or Sez6, protein is located in developing dendrites and is also present in dendritic spines, specialized structures on dendrites that receive and integrate excitatory signals. Furthermore, when this protein is lacking, dendrites and excitatory synapses fail to form correctly4. The probable dominant isoform of the protein has features of a transmembrane receptor although, when the subcellular distribution of immunolabeled protein was examined by confocal microscopy or by immunoelectron microscopy most, if not all, of the signal appeared associated with small vesicles in the somatodendritic compartment with little, or no, protein labeled on the plasma membrane at the cell surface.
In order to definitively show that this putative receptor with a predicted large extracellular domain is trafficked to the plasma membrane, we adopted a live-cell approach using the antiserum we had generated to an extracellular portion of the protein to label protein on the cell surface. By combining this &#8220;antibody feeding&#8221; approach with two applications of differentially-labeled secondary antibody separated by an extensive blocking step and a permeabilization step, we were able to identify two different pools of protein distinguished by binding to fluorescently-labeled secondary antibodies bearing different fluorescent tags. Thus, we were able to distinguish protein that had been internalized by endocytosis during the antibody incubation step from protein that either remained on the cell surface or was trafficked to the surface during this period. Using this method, we established that the protein of interest is trafficked to and from the cell surface in neurons. Therefore, this relatively fast and simple technique proved more informative than traditional immunocytochemistry methods or pre-embedding immunogold electron microscopy, despite the fact that we used the same rabbit polyclonal antiserum for all these techniques. This technique is generally applicable to any transmembrane protein provided a good antibody recognizing extracellular domain epitopes is available. The technique has been used previously to study receptor trafficking of the glutamate receptor GluR1 subunit5.

<table border="0" cellpadding="0" cellspacing="0" width="937">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">PBS with Ca and Mg</td>
			<td style="width:269px;">Invitrogen</td>
			<td style="width:119px;">14040182</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Neurobasal medium</td>
			<td style="width:269px;">Invitrogen</td>
			<td style="width:119px;">21103-049</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">B27 supplement</td>
			<td style="width:269px;">Invitrogen</td>
			<td style="width:119px;">17504-044</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">L-glutamine</td>
			<td style="width:269px;">Invitrogen</td>
			<td style="width:119px;">25030-081</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Papain Dissociation System</td>
			<td style="width:269px;">Worthington Biochemical Corporation</td>
			<td style="width:119px;">PDS</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Bovine Serum Albumin&#160;</td>
			<td style="width:269px;">Sigma Aldrich Australia</td>
			<td style="width:119px;">A9418</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:383px;">Hank&#39;s balanced salt solution without calcium, magnesium, phenol red</td>
			<td style="width:269px;">Invitrogen (Gibco)</td>
			<td style="width:119px;">14175-079</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Poly-D-Lysine</td>
			<td style="width:269px;">Sigma Aldrich Australia</td>
			<td style="width:119px;">P0899</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Natural mouse laminin</td>
			<td style="width:269px;">Invitrogen</td>
			<td style="width:119px;">23017-015</td>
			<td>Thaw on ice prior to making aliquots</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Fetal bovine serum HyClone</td>
			<td style="width:269px;">Thermo Fisher</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">fluorodeoxyuridine</td>
			<td style="width:269px;">Sigma Aldrich Australia</td>
			<td style="width:119px;">F0503</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">uridine</td>
			<td style="width:269px;">Sigma Aldrich Australia</td>
			<td style="width:119px;">U3003</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">18 mm round glass coverslips</td>
			<td style="width:269px;">Menzel Gl&#228;ser</td>
			<td>CB00180RA1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">VECTASHIELD aqueous mounting medium</td>
			<td style="width:269px;">Vector Laboratories</td>
			<td style="width:119px;">H1400</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Donkey anti-rabbit Dylight 649&#160;</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td style="width:119px;">711-495-152</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">AffiniPure Fab fragment Goat anti-Rabbit 1gG (H+L)&#160;</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td>111-007-003</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Paraformaldehyde</td>
			<td style="width:269px;">Sigma Aldrich Australia</td>
			<td style="width:119px;">P6148</td>
			<td>TOXIC - handle in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:383px;">Triton-X-100</td>
			<td style="width:269px;">Sigma Aldrich Australia</td>
			<td style="width:119px;">T8787</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexafluor 488-conjugated donkey anti-rabbit 2&#176; antibody</td>
			<td style="width:269px;">Invitrogen - Molecular Probes</td>
			<td style="width:119px;">A21206</td>
			<td></td>
		</tr>
	</tbody>
</table>

differential labeling of cell-surface and internalized proteins after antibody feeding of live cultured neurons

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of live neurons with a specific primary antibody raised against an extracellular portion of the protein. Given that receptors are trafficked to and from the surface, if cells are permeabilized after fixation then both cell-surface and internal protein will be detected by the same labeled secondary antibody. Here, we adapted a method used to study protein trafficking (&#8220;antibody feeding&#8221;) to differentially label protein that had been internalized by endocytosis during the antibody incubation step and protein that either remained on the cell surface or was trafficked to the surface during this period. The ability to distinguish these two pools of protein was made possible through the incorporation of an overnight blocking step with highly-concentrated unlabeled secondary antibody after an initial incubation of unpermeabilized neurons with a fluorescently-labeled secondary antibody. After the blocking step, permeabilization of the neurons allowed detection of the internalized pool with a fluorescent secondary antibody labeled with a different fluorophore. Using this technique we were able to obtain important information about the subcellular location of this putative receptor, revealing that it was, indeed, trafficked to the cell-surface in neurons. This technique is broadly applicable to a range of cell types and cell-surface proteins, providing a suitable antibody to an extracellular epitope is available.

The dual-color fluorescent immunostaining technique presented here is useful for labeling extracellular domains of transmembrane proteins in living cells (shown schematically in Figure&#160;1). During the incubation period, the immunoglobulins bind accessible epitopes and a proportion of the population of protein molecules, together with bound antibody, is endocytosed. In addition, newly synthesized protein may reach the cell surface via forward trafficking and recycled protein molecules may be returned ...

Watch this Scientific Journal Video about Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons at JoVE.com

Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

We describe a method to label protein on the surface of living neurons using a specific polyclonal antibody to extracellular epitopes. Protein bound by the antibody on the cell surface and subsequently internalized via endocytosis can be distinguished from protein remaining on, or trafficked to, the surface during the incubation.

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of ...

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Research

JoVE Journal

Neuroscience

15.4K vistas.  The University of Melbourne.  Se describe un método para etiquetar proteína en la superficie de neuronas vivas usando un anticuerpo policlonal específico para epítopos extracelulares. La proteína unida por el anticuerpo en la superficie celular y, posteriormente, internalizado a través de la endocitosis se puede distinguir de la proteína restante en, o de trata a, la superficie durante la incubación. 

Video: Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.