We are grateful to the Wellcome Trust and the ERC for financial support. IMGG is an EMBO Fellow. KLH is a BBSRC-funded PhD student. We thank Philip Rubin and Patrick Tidball for technical and cell culture support and Dr Andrew Doherty for maintenance and assistance with the microscopes.

1. Cell Culture, Viral Transduction, and Protein Expression
<ol>
 <li>Culture high density hippocampal neurons from embryonic day 18 (E18) rat pups on poly-l-lysine-coated glass coverslips for 14-25 days in vitro (DIV).</li>
 <li>6-24hrs prior to the live experiments, transduce cells with attenuated Sindbis virus containing the membrane protein of interest, tagged with the super-ecliptic pHluorin (SEP).</li>
 <li>Add the pseudovirion-containing medium directly to the coverslip containing 1mL of conditioned media and returned to the culture incubator. The titer and time for protein expression following viral transduction will vary depending on the virus batch and should be determined for each batch prior to commencing live cell experiments.</li>
</ol>
2. FRAP-FLIP Live Cell Imaging
<ol>
 <li>Equipment set-up
 <ol>
 <li>Transfer the coverslip to the imaging chamber of a Zeiss Axiovert LSM 510 META confocal microscope. To minimize power fluctuations during imaging, ensure the microscope has been switched on, with 100% laser output, for at least 20 mins prior to imaging.</li>
 <li>Immediately, replace the culture medium with pre-warmed (37 &deg;C) extracellular recording solution containing 140 mM NaCl, 5 mM KCl, 15 mM glucose ,1.5 mM CaCl2, 1.5 mM MgCl2, 20-25 mM HEPES (pH adjusted to 7.4 with NaOH) and place the chamber on the pre-heated stage (37 &deg;C) of the Zeiss Axiovert. 
 
 Ensure the osmolarity of the extracellular recording solution is adjusted to within 10 mOsM of your culture medium. Provided no significant evaporation occurs during the timecourse of the experiment, this CO2 independent solution is suitable for short experiments (&lt;10 hrs). Supplementing with 1-2 mM sodium bicarbonate is recommended.</li>
 </ol>
 </li>
 <li>Defining the imaging capture parameters
 <ol>
 <li>First, identify a neuron expressing the recombinant protein of interest and bring it into focus.</li>
 <li>With a 63X oil objected, acquire an image of the whole cell using 488nm laser light excitation at low laser power. To minimize photobleaching, use a fast nominal speed (7-9) and low pixel resolution (512-512) keeping total scan speed &lt;1 second.</li>
 <li>Select a portion of dendrite to image and zoom to capture a frame containing the ROI (~1.5-2.5 x optical zoom). Where possible, ensure the field of view contains several processes so that measurements from reference dendrites can be obtained, to determine if non-specific photobleaching due to acquisition is occurring.</li>
 <li>Adjust the filters, pinhole, scan speed and detector gain to enable maximal fluorescence from minimal laser excitation but with limited saturation. A large pinhole diameter is recommended, to maximize photon collection (2&mu;m is suitable for spines and ternary dendrites). The detector gain should be strong enough to detect small fluorescence increments, such that the very first images, before the photobleaching do not overcome 10% of saturated pixels.</li>
 <li>Save this configuration to be used for the pre-/ post-bleach and recovery phases of the experiment.</li>
 <li>Next define the photobleach regions; selecting an ROI for the initial photobleach and flanking regions for the subsequent repetitive photobleaching phase. Ensure the flanking regions are wide enough to prevent recovery by diffusion between scans (typically 5 &mu;m, see Fig. 2).</li>
 <li>Adjust the bleach parameters for both photobleach ROIs. The initial photobleach should be rapid (0.1-0.5 sec) requiring between 1-5 iterations, depending on the optical zoom and the volume of the ROI. For the flanking regions, the laser excitation should be adjusted to ensure continuous photobleaching of the flanking regions, but without phototoxic damage.</li>
 <li>As a guideline, we use 100% laser excitation for the initial photobleach, and 10% for the repetitive photobleaching phase.</li>
 </ol>
 </li>
 <li>Image acquisition
 <ol>
 <li>Once all parameters have been set, perform the FRAP-FLIP experiment as a variable time-lapse image sequence, outlined in the 4 blocks below: 
 
 BLOCK 1: 3-10 pre-bleach baseline images, at low laser power, no time delay 
 BLOCK 2: Photobleach the central ROI at full laser power, 1-5 iterations 
 BLOCK 3: 3-10 post-bleach recovery images 
 BLOCK 4: Repetitive photobleach of flanking regions at medium laser power with image capture at a typical time interval of 1 - 5 seconds, depending on the recovery rate of the protein under investigation.</li>
 <li>Finally, replace the extracellular recording solution with recording solution buffered at pH6 containing 140 mM NaCl, 5 mM KCl, 15mM glucose ,1.5 mM CaCl2, 1.5 mM MgCl2, 20-25 mM MES (to quench fluorescence) or complemented with 50 mM NH4Cl containing 90 mM NaCl, 5 mM KCl, 15 mM glucose ,1.8 mM CaCl2, 0.8 mM MgCl2, 20-25 mM HEPES, pH7.4 (to reveal the proteins in low pH intracellular stores), (see Fig. 2).</li>
 <li>Collect at least 10 to 20 data sets for each recombinant protein, to enable statistical analysis. To avoid bias results, ensure imaging conditions are maintained consistent across replicates. Discard any data sets where incomplete bleaching, significant focal plane drift or phototoxic cell damage are observed.</li>
 </ol>
 </li>
</ol>
3. Data Analysis
<ol>
 <li>Open the images with ImageJ software.</li>
 <li>Align the stacks to account for small fluctuations in the xy plane that may have occurred throughout the time series using the Stackreg plugin (plugins &rarr; stackreg &rarr;transformation: rigid body &rarr; &lt;ok&gt; ). </li>
 <li>For images taken on Zeiss confocal, use the LSM Toolbox plugin to report the time values as a text file (plugins&rarr; LSM Toolbox &rarr; Show LSM Toolbox &rarr; &lt;apply stamps&gt; &rarr; &lt;apply t-stamp&gt; ), and import these values into an analysis spreadsheet. </li>
 <li>To measure the fluorescence fluctuations during the FRAP-FLIP experiment, divide the photobleached segment into individual pixel regions (~20pixels) and measure the mean fluorescence of these ROIs at each time point (F). To quickly obtain these values, select multiple ROIs using the ImageJ 'ROI Manager' analysis tool (Analyze &rarr; Tools &rarr; ROI Manager) and report the mean fluorescence per pixel using the 'Plot Z-axis profile' command ( Image &rarr;Stacks &rarr;Plot Z-axis profile).</li>
 <li>Repeat this step to measure the fluorescence intensity of a background, non fluorescent region.</li>
 <li>Normalize the fluorescence intensity at each time point by subtracting the background values to remove experimental noise, and divide all values by the mean pre-bleached baseline measure.</li>
 <li>Measure the fluorescence intensity of adjacent non-photobleached dendrites to assess the level of non-specific photobleaching during acquisition. Correction for non-specific photobleaching is discouraged for interpretation of 'FRAP-FLIP' data and should be avoided whenever possible.</li>
 <li>To calculate if significant recovery has taken place in a ROI, subtract the mean of last 5-10 measures of the sequence from the mean of the first 5-10 measures (&Delta;F) and determine the statistical significance. Categorize the recovering and non-recovering ROIs to assess the pattern of exocytosis (i.e. exocytosis hotspots or spine vs shaft recovery).</li>
</ol>
4. Representative Results
The outcome of typical FRAP-FLIP experiment is shown in Fig. 2. Here, a neuron expressing the GluA2 subunit of AMPA type glutamate receptor, tagged with SEP has been selectively photobleached along a region of dendrite. Fig 2.A illustrates the region of dendrite which has been imaged and indicates the ROIs that have been selected for photobleaching (white box regions). The large white boxed area was bleached once, followed by repetitive bleaching of the flanking boxed regions, highlighted with double headed blue arrows. The red arrow indicates the measured ROI, shown in high magnification in the lower panels.
Fig 2.B, shows the fluorescence intensity in the measured ROI over the time course of the experiment. In this example, a one minute recovery period was recorded after the initial photobleach to allow unbleached receptors to enter the ROI by lateral diffusion. When the flanking regions are photobleached, the fluorescence signal from this highly motile fraction of receptors is occluded from the central region, and those within the region are diluted out. The increase in fluorescence (&Delta;F) observed over the 'FLIP' sequence can therefore be attributed to insertion of SEP-GluA2 in the dendritic shaft. The low pH wash and pH 7.4 + NH4Cl addition respectively confirm that the measured fluorescence is derived from surface proteins and reveal the proportion of intracellular, sequestered proteins within the measured ROI.
In contrast to FRAP, this methodology isolates recovery due to exocytosis, resulting in a greatly reduced level of fluorescence recovery in the photobleached region. To date no reliable mathematical model has been developed to fit and analyze the recovery traces recorded using this selective bleaching technique. It is however, possible to fit the recovery trace with a mono exponential recovery:
F(t)=As - A0e(-t/&tau;)
Where F(t) is fluorescence at time t, As is steady state value, A0 is the offset at time 0, &tau; is the time constant. The recovery growth is fixed with a given rate to reach an equilibrium, corresponding to a steady state between insertion and diffusion. Importantly, the time constant extracted from this analysis does not reflect the time constant of exocytosis and can only be used for comparative treatments for an individual protein.
Moreover, the expected pattern of fluorescence recovery is likely to be observed in sub-domains along the photobleached region. Analysis of small ROI within a photobleached segment may be essential to reveal exocytosis "hotspots" and it is advisable to analyse regions of comparable lengths as the variability of density of these spots among the dendrite will affect the calculated rate.
Fig 3 shows an extension of this protocol, applying the photobleach to large section ROI with multiple dendrites in the field of view, followed by selective repetitive bleaching of only one dendrite. This approach enables traditional 'FRAP' and 'FRAP-FLIP' data to be acquired in parallel. In this example, hippocampal neurons have been infected with a glutamate receptor subunit of the kainate class; SEP-tagged GluK2. Prior to imaging, these neurons -were treated with cylcohexamide (2hrs at 200&mu;g/ml), to block protein synthesis. As such, fluorescence recovery in the respective measured ROIs (Fig 3.B) reveals the proportion of recovery due to lateral diffusion and recycling in the standard FRAP dendrite versus recycling alone in the dendrite subjected to the 'FRAP-FLIP' protocol. Comparing the &Delta;F values from the FRAP versus 'FRAP-FLIP' recovery curves, the relative contributions of recycling (&Delta;Frec = 11.05%) versus lateral diffusion (&Delta;Fdiff = 9.35%) can be inferred. Unlike Figure 2, the recovery does not maintain a steady state level but rather, due to the inhibition of protein synthesis, shows a transient increase and subsequent drop-off in signal, corresponding to depletion of the available receptor pool (approximately 20% decline observed over the recording period).
<img alt="Figure 1" src="/files/ftp_upload/3747/3747fig1.jpg" />
Figure 1. Schematic of the principals of FRAP vs FRAP-FLIP protocols This schematic illustrates the outcomes of a regular FRAP versus a 'FRAP-FLIP' protocol, using SEP-tagged receptors. Fluorescence recovery in traditional FRAP is shown on left-hand side. Measured fluorescence recovery in the central ROI is attributed to a combination of lateral diffusion f non-photobleached SEP-tagged receptors from outside the photobleached ROI and insertion of receptors through recycling and/or de novo exocytosis into the dendritic shaft. By contrast, the repetitive photobleached of the flanking ROIs illustrated in right-hand side, shows how this modified 'FRAP-FLIP' protocol silences recovery due to lateral diffusion. As such, any measured fluorescence recovery can be attributed to direct insertion in the ROI.
<img alt="Figure 2" src="/files/ftp_upload/3747/3747fig2.jpg" />
Figure 2. Insertion of SEP-GluA2 into the plasma membrane on the dendritic shaft
A) A hippocampal neuron expressing SEP-GluA2, selectively photobleached along a region of dendrite. The schematic illustrates the region of dendrite which has been imaged, while the upper left hand panel highlights the ROIs that have been selected for photobleaching. The large white boxed area was bleached once, followed by repetitive bleaching of the flanking boxed regions, highlighted with double headed blue arrows. This panel shows the dendrite prior to photobleaching. The red arrow indicates the measured ROI, shown in high magnification in the lower panels. B) shows the fluorescence intensity in the measured ROI over the time course of the experiment, plotted as gray level in the ROI (not normalized). The black arrow highlight the timepoint of the initial photobleach and green arrow indicates the start of the repetitive photobleaching. &Delta;F indicates the increase in fluorescence observed over the recovery period, due to insertion of SEP-GluA2 into the dendritic shaft. Subsequent low pH and pH 7.4 + NH4Cl washes confirm that the recovered fluorescence relates to surface expressed receptors.
<img alt="Figure 3" src="/files/ftp_upload/3747/3747fig3.jpg" />
Figure 3. Recycling &amp; lateral diffusion vs Recycling of SEP-GluK2 on the dendritic shaft following cyclohexamide treatment
A) A hippocampal neuron expressing SEP-GluK2, selectively photobleached with parallel FRAP and 'FRAP-FLIP' recovery protocols performed along separate dendritic ROIs. This neuron was subject to pretreatment with cyclohexamide, to block protein synthesis (2 hrs at 200 &mu;g/ml). The schematic illustrates the region of dendrite which has been imaged, while the left hand panel highlighting the ROIs that have been selected for photobleaching. The large white boxed area was bleached once, followed by repetitive bleaching of the flanking boxed regions, of the lower dendrite only, highlighted with double headed blue arrows. This panel shows the dendrites prior to photobleaching. Red arrows highlight the ROIs shown in high magnification in B) illustrating the fluorescence recovery in traditional FRAP versus the 'FRAP-FLIP' protocol. Comparisons of levels of fluorescence intensity in the late stage of recovery (third panels) show the contribution of lateral diffusion and recycling versus recycling alone. C) shows the fluorescence intensity in the measured ROIs over the time course of the experiment, plotted as normalized intensity values. The black arrow highlight the timepoint of the initial photobleach and green arrow indicates the start of the repetitive photobleaching. &Delta;Frec indicates the transient recovery due to receptor recycling, determined from the FRAP/FLIP dendrite while &Delta;F diff measures the contribution of lateral diffusion of SEP-GluK2 into the dendritic shaft in the 'FRAP only' ROI. The transient increase is followed by gradual decline in signal, corresponding to the run down of available receptors as a result of cyclohexamide treatment. The subsequent low pH and pH 7.4 + NH4Cl washes confirm that the recovered fluorescence relates to surface expressed receptors.

<ol>
	<li>Sankaranarayanan, S., De Angelis, D., Rothman, J. E., Ryan, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+pHluorins+for+optical+measurements+of+presynaptic+activity.">The use of pHluorins for optical measurements of presynaptic activity.</a> Biophys. J. 79, 2199-2208 (2000).</li><li>Ashby, M. C., Ibaraki, K., Henley, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=It's+green+outside:+tracking+cell+surface+proteins+with+pH-sensitive+GFP.">It's green outside: tracking cell surface proteins with pH-sensitive GFP.</a> Trends Neurosci. 27, 257-261 (2004).</li><li>Swaminathan, R., Hoang, C. P., Verkman, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photobleaching+recovery+and+anisotropy+decay+of+green+fluorescent+protein+GFP-S65T+in+solution+and+cells:+cytoplasmic+viscosity+probed+by+green+fluorescent+protein+translational+and+rotational+diffusion.">Photobleaching recovery and anisotropy decay of green fluorescent protein GFP-S65T in solution and cells: cytoplasmic viscosity probed by green fluorescent protein translational and rotational diffusion.</a> Biophys. J. 72, 1900-1907 (1997).</li><li>Patterson, G. H., Knobel, S. M., Sharif, W. D., Kain, S. R., Piston, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+the+green+fluorescent+protein+and+its+mutants+in+quantitative+fluorescence+microscopy.">Use of the green fluorescent protein and its mutants in quantitative fluorescence microscopy.</a> Biophys. J. 73, 2782-2790 (1997).</li><li>Wiedenmann, J., Oswald, F., Nienhaus, G. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+proteins+for+live+cell+imaging:+opportunities,+limitations,+and+challenges.">Fluorescent proteins for live cell imaging: opportunities, limitations, and challenges.</a> IUBMB Life. 61, 1029-1042 (2009).</li><li>Ashby, 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=Removal+of+AMPA+receptors+(AMPARs)+from+synapses+is+preceded+by+transient+endocytosis+of+extrasynaptic+AMPARs.">Removal of AMPA receptors (AMPARs) from synapses is preceded by transient endocytosis of extrasynaptic AMPARs.</a> J. Neurosci. 24, 5172-5176 (2004).</li><li>Bouschet, T., Martin, S., Henley, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Receptor-activity-modifying+proteins+are+required+for+forward+trafficking+of+the+calcium-sensing+receptor+to+the+plasma+membrane.">Receptor-activity-modifying proteins are required for forward trafficking of the calcium-sensing receptor to the plasma membrane.</a> J. Cell. Sci. 118, 4709-4720 (2005).</li><li>Ashby, M. C., Maier, S. R., Nishimune, A., Henley, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lateral+diffusion+drives+constitutive+exchange+of+AMPA+receptors+at+dendritic+spines+and+is+regulated+by+spine+morphology.">Lateral diffusion drives constitutive exchange of AMPA receptors at dendritic spines and is regulated by spine morphology.</a> J. Neurosci. 26, 7046-7055 (2006).</li><li>Martin, S., Bouschet, T., Jenkins, E. L., Nishimune, A., Henley, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bidirectional+regulation+of+kainate+receptor+surface+expression+in+hippocampal+neurons.">Bidirectional regulation of kainate receptor surface expression in hippocampal neurons.</a> J. Biol. Chem. 283, 36435-36440 (2008).</li><li>Jaskolski, F., Mayo-Martin, B., Jane, D., Henley, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamin-dependent+membrane+drift+recruits+AMPA+receptors+to+dendritic+spines.">Dynamin-dependent membrane drift recruits AMPA receptors to dendritic spines.</a> J. Biol. Chem. 284, 12491-12503 (2009).</li></ol>

No conflicts of interest declared.

We describe an innovative strategy to visualize the components of plasma membrane protein trafficking. The combinatorial approach of photobleaching techniques with SEP-tagged protein enables selectively plasma membrane insertion events to be assessed. By continually photobleaching the membrane proteins in flanking regions during recovery, the 'FRAP-FLIP' method assesses the contribution of vesicular trafficking to fluorescence recovery. This novel approach allows direct recording of protein membrane insertion, enabling both the number of sub-compartments where recovery is observed and the amplitude of the recovery (&Delta;F) at the steady state to be determined. Further, comparison of FRAP with and without FLIP allows the proportion of recovery attributable to lateral diffusion to be calculated.
Moreover, in the same experiment, regions of non-photobleached dendrite adjacent to the flanking photobleached regions can be qualitatively assessed during the recovery; fluorescence loss observed in these regions will be due to lateral diffusion of photobleached receptors into these non-photobleached segments of the dendrite.
 This selective photobleaching protocol can be utilized to investigate a variety of cellular trafficking processes such as characterizing excocytosis in the plasma membrane in defined subareas (for example, dendrites or spines) or assessing the contribution of lateral diffusion vs insertion by conducting parallel FRAP and 'FRAP-FLIP' protocols along adjacent dendrites (Fig. 3). 
 Clearly, while specific guidelines have been presented, each lab will need optimize the imaging parameters according to specific specimens and equipments. Importantly, all surface receptors in the ROI need to be photobleached, independent of the z-axis focal plane, but without significant phototoxic damage or non-specific photobleaching. Users should exercise caution when attempting this protocol at the cell soma, as the high percentage of relatively low pH intracellular compartments typically results in high background fluorescence in these regions. Moreover, whilst we have demonstrated how this technique can be applied in varies ways, prior to commencing selective photobleaching experiments on a new SEP-tagged construct, we advise that an initial characterization of the tagged receptors is first conducted, as described by Ashby et al2. 
Overall, this method is a powerful and versatile adaptation of the standard FRAP protocol, enabling insertion events in plasma membrane to be evaluated in near real time.

<table border="1">
	<tbody>
 	<tr>
 	<td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 	<td>24mm glass coverslips</td>
 <td>VWR International</td>
 <td>631-0161</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Poly-l-lysine</td>
 <td>Sigma</td>
 <td>P2636</td>
 <td>1mg/ml in borate buffer to coat coverslips</td>
 </tr>
 <tr>
 	<td>Neurobasal Medium</td>
 <td>Invitrogen</td>
 <td>21103</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>B27</td>
 <td>Invitrogen</td>
 <td>17504-044</td>
 <td>2% in neuronal plating and feeding medium</td>
 </tr>
 <tr>
 	<td>Penicillin Streptomycin</td>
 <td>Sigma</td>
 <td>P0781</td>
 <td>1% in neuronal plating and feeding medium</td>
 </tr>
 <tr>
 	<td>L-Glutamine</td>
 <td>Invitrogen</td>
 <td>25030</td>
 <td>2mM / 0.8mM in plating/feeding medium</td>
 </tr>
 <tr>
 	<td>Horse Serum</td>
 <td>Biosera</td>
 <td>DH-291H</td>
 <td>10% in neuronal plating media only</td>
 </tr>
 <tr>
 	<td>pSIN ReP5 cloning vector</td>
 <td>Invitrogen</td>
 <td>K75001</td>
 <td>For Sindbis virus production</td>
 </tr>
 <tr>
 	<td>LSM510 META confocal system</td>
 <td>Zeiss</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Image J Software</td>
 <td>NIH</td>
 <td>&nbsp;</td>
 <td>open access software. All plugins described herein are available at <a href="http://rsbweb.nih.gov/ij/plugins/">http://rsbweb.nih.gov/ij/plugins/</a></td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching at JoVE.co

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

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 ...

lateral-diffusion-exocytosis-membrane-proteins-cultured-neurons

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JoVE Journal

Neuroscience

83.0K 次观看.  University of Bristol. 这份报告介绍了利用活细胞成像和光漂白技术，以确定表面的表达，运输途径和贩运外生表示，pH敏感的绿色荧光蛋白标记的蛋白质在神经元细胞膜动力学。

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

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 ...

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

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 ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

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 ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

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.

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 ...

Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient

Neuronal subcellular fractionation techniques allow the quantification of proteins that are trafficked to and from the synapse. As originally described in the late 1960&#8217;s, proteins associated with the synaptic plasma membrane can be isolated by ultracentrifugation on a sucrose density gradient. Once synaptic membranes are isolated, the macromolecular complex known as the post-synaptic density can be subsequently isolated due to its detergent insolubility. The techniques used to isolate synaptic plasma membranes and post-synaptic density proteins remain essentially the same after 40 years, and are widely used in current neuroscience research. This article details the fractionation of proteins associated with the synaptic plasma membrane and post-synaptic density using a discontinuous sucrose gradient. Resulting protein preparations are suitable for western blotting or 2D DIGE analysis.

This article details the enrichment of proteins associated with the synaptic plasma membrane by ultracentrifugation on a discontinuous sucrose gradient. The subsequent preparation of post-synaptic density proteins is also described. Protein preparations are suitable for western blotting or 2D DIGE analysis.

Neuronal subcellular fractionation techniques allow the quantification of proteins that are trafficked to and from the synapse. As originally described in ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

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, ...