The authors would like to thank Drs. C. Szalinski, H. Teng, and M. Bruchez for help with Imaris, R. Vistein, and D. Shiwarski for technical help and advice, and Dr. M von Zastrow, Dr. T Kirchhausen, Dr. D Drubin, and Dr. W Almers for reagents and helpful discussion. Funding provided by T32 grant NS007433 to SLB and NIH DA024698 and DA036086 to MAP.

1. Expression of Fluorescently Tagged CME Components in Cultured Cells
HEK293 cells are useful model cells that have been used extensively to study GPCR biology and endocytosis, and therefore are used as models in this protocol. Use any transfection protocol providing uniform expression without overexpression and low cytotoxicity.
<ol>
	<li>Handle all cell culture in a sterile laminar flow hood. Fill 4 wells of a 12-well plate with 1 ml of Dulbecco&#8217;s Modified Essential Media (DMEM) with 10% Fetal Bovine Serum (FBS). From a confluent T25 flask of HEK293 cells dilute 1:50, 1:25, 1:16, 1:10 respectively into each of the 4 wells.
	<ol>
		<li>Alternatively, seed cells to a multi-well plate of desired size, as per manufacturer&#8217;s instructions. Seed 0.4 - 2 x 104 cells into a 12-well plate. Grow the cells overnight in a sterile 37 &#176;C with 5% CO2 and 95% humidity. 
		NOTE: Considering that growth rates vary depending on the conditions and cell lines, it might be preferable to seed multiple wells to ensure ideal confluency for transfection the next day or the possibility of performing transfections in duplicate.</li>
	</ol>
	</li>
	<li>The next morning, select a well that is ~70% confluent for transfection (see Figure 1B). If no well has reached this state, wait another day. 
	NOTE: 70% confluency is ideal for HEK293 cells. Transfecting cells that are more sparse results in cell death and toxicity. Transfecting over-confluent cells results in poor expression.</li>
	<li>Add 60 &#181;l buffer EC (from transfection kit), 400 ng of plasmids encoding tagged GPCRs and/or components of the clathrin machinery, and 2.4 &#181;l of Enhancer into the 1.5 ml microcentrifuge tube, as described in the manufacturer&#8217;s protocol. Vortex for 2 sec to ensure sufficient mixing, and spin in a microfuge at 2,000 x g for 2 sec to collect the liquid at the bottom.</li>
	<li>Add 6 &#956;l Effectene as described in the manufacturer&#8217;s protocol. Vortex for 2 sec, and spin at 2,000 x g for 2 sec in a microfuge to collect the liquid at the bottom. Wait for 20 min to allow for formation of transfection complex micelles.</li>
	<li>Aspirate media from the well to be transfected. Gently add 0.8 ml of DMEM with 10% FBS to the transfection reagent mixture and mix by pipetting up and down slowly to reduce breaking the micelles. Add the media and transfection mixture to the well very slowly from the side. Add any remaining transfection mixture from the microcentrifuge tube to the well using a micropipettor. 
	NOTE: Optionally, add an extra 0.5 ml of DMEM with 10% FBS to the transfected well to give the cells extra nourishment resulting in gentler transfection and lower transient expression.</li>
	<li>Return the cells to the 37 &#176;C incubator for 5 hr.</li>
	<li>Aspirate media containing the transfection mixture. Replace with 1 ml of DMEM with 10% FBS. Allow the cells to grow overnight.</li>
	<li>The next day, pass the cells to uncoated coverslips, previously sterilized by autoclaving.
	<ol>
		<li>Add 0.5 ml of Phosphate Buffered Saline (PBS) with 1 mM EDTA to each well. Alternatively, use 0.5 ml of 0.05% Trypsin-EDTA. Leave in the incubator for 1 min.</li>
		<li>Place 3 round, sterile 25 mm coverslips into separate wells of a 6-well plate. Fill each well with 2 ml of media. Gently push the coverslip down to the bottom of the well to prevent cells from growing on the bottom of the coverslip.</li>
		<li>Manually agitate the well to lift off the cells from the bottom of the well. Add 1 ml of DMEM with 10% FBS to resuspend. Distribute the lifted cells from 1 well of a 12-well plate evenly among the 3 coverslips. Alternatively, plate cells on 3 glass-bottomed dishes.</li>
	</ol>
	</li>
	<li>Allow the cells to grow on the coverslips for at least 48 hr before imaging. Ensure that cells are spread out and flat.</li>
</ol>
2. Imaging CCP and Cargo Dynamics Using TIRFM
<ol>
	<li>Conjugate M1 anti-FLAG antibodies with Alexa dyes using Alexa Fluor Protein Labeling kit according to the manufacture&#8217;s protocol. Alternatively, pre-conjugated antibodies may be purchased.</li>
	<li>Pre-incubate cells with the antibodies at a 1:1,000 dilution in 1 ml of pre-warmed Leibovitz medium with 10% FBS, or Opti-MEM with 50 mM HEPES pH 7.4 and 10% FBS for 10 min at 37 &#176;C (imaging medium), to label FLAG-tagged GPCRs on the cell surface. Skip these steps if genetically encoded fluorescent proteins, such as GFP, are used as tags (see discussion for details).</li>
	<li>Transfer the coverslip to a live-cell imaging chamber. Alternatively, for glass-bottomed dishes, aspirate antibody-labeling media and add 700 &#181;l of pre-warmed imaging medium.
	<ol>
		<li>Use a pair of forceps and bend the tip of a 25 gauge needle into a hook.</li>
		<li>Hold a pair of forceps in the dominant hand. Hold the bent needle in the other. Turn the needle until the hook curves down towards the glass. Gently drag the needle, hook side down, across the bottom of a 6-well plate until it hooks onto the coverslip. Take care to not scratch the surface of the coverslip, as it will detach cells. Gently lift the coverslip up from the bottom of the well. Balance the coverslip against wall of the well for support.</li>
		<li>Use the forceps to grasp near the edge of the coverslip and move the coverslip cell side up to the imaging chamber, and assemble the chamber. Add 700 &#181;l (or appropriate amount) of pre-warmed imaging medium. Handle the coverslips carefully without pressure to prevent cracking or breaking.</li>
	</ol>
	</li>
	<li>Transfer the chamber onto the microscope and keep the temperature of the cells at 37 &#176;C using a stage heater or an enclosed incubator.</li>
	<li>Bring cells into focus using a 100X or 60X TIRF oil-immersion objective (1.45 NA or above). Image the cells in conventional epifluorescence or confocal modes of illumination (i.e. not TIRFM) to identify cells expressing the appropriate constructs. Out-of-focus cells are nearly invisible in the thin depth of TIRF illumination making it difficult to find the focal plane and cells. 
	NOTE: Some systems use the same lasers for TIRFM to image in conventional epifluorescence, by moving the angle of incidence of the laser above the critical angle. As an important safety precaution, always be aware of the angle of the laser beam, and always direct it away from the observer.</li>
	<li>Focus down to the bottom surface of an expressing cell until the outline of plasma membrane around the cell disappears and the center of the cell appears. This is most apparent with a plasma membrane localized cargo protein such as the &#181;-opioid receptor (see Figure 2A).</li>
	<li>Switch to TIRF imaging.
	<ol>
		<li>If using the same lasers for imaging in conventional epifluorescence, increase the angle of incidence of the laser until out-of-focus fluorescence in the interior of the cell disappears, and no outline of plasma membrane around the cell is seen. (Compare Figure 2C to Figures 2A and 2B) 
		NOTE: In TIRFM, moving the focal plane causes the image to go completely out of focus and/or dim, and the overall fluorescence levels decreases due to the smaller depth of illumination. Therefore, an alternate method to switch to TIRFM illumination is to first focus on the center of the cell in confocal/conventional epifluorescence, then increase the angle of incidence until you lose most of the fluorescence, and then focus down to the plasma membrane.</li>
		<li>Once in TIRFM, ensure the excitation laser reflects back through the objective and is not visible above the objective. Confirm this by checking if the laser spot is visible. 
		NOTE: In conventional epifluorescence or confocal modes, the laser illumination is visible above the objective, such as on the ceiling.</li>
		<li>Refine the focus to get a crisp image. The main components of the endocytic machinery, such as clathrin, will be visible in puncta. Adjust the focus on fine, so the puncta are as round as possible. 
		NOTE: For membrane-localized cargo, adjust the fine focus until the filopodia appear distinct.</li>
		<li>Use an acquisition program to save all the imaging parameters such as mirrors and TIRF angle. Do this for all necessary wavelengths.</li>
	</ol>
	</li>
	<li>Scan around the coverslip to find cells expressing all desired endocytic markers: the &#181;-opioid receptor and the adapter &#946;-arrestin in the example shown in Figure 3A. Select cells that are expressing but not over-expressing. See discussion for details.</li>
	<li>Once cells are identified, acquire images every 3 sec for 10 min. This is sufficient to capture the lifetime of a CCP, since the average lifetime of a CCP in HEK293 cells imaged under these conditions is around 40 sec10-11. This allows about 12-14 frames to observe the CCP. For example see Movie 1. Repeat all steps to image additional cells.</li>
</ol>
3. Analysis of Endocytic Dynamics by Manual Verification
Although manual verification of CCP lifetimes is greatly limited by the number of CCPs that can be detected, it still remains an accurate means of detection undeterred by global changes in the image and detection artifacts. See discussion for details.
<ol>
	<li>Open the file in ImageJ. Images are stored as a single stack of tiff files with interleaved channels. Convert images to hyperstacks. Go to Image &#62; Hyperstacks &#62; Stack to Hyperstack. Enter the number of channels acquired (i.e. if imaging &#946;-arrestin and the &#181;-opioid receptor, enter 2), Enter 1 for Z stack (TIRFM is a single plane), and the number of frames of the movie (i.e. a 10 min movie at 1 frame every 3 sec is 200) in the pop-up window.</li>
	<li>The hyperstack format yields a window with two scroll bars. Use the top bar to move through the channels and the bottom to move through time. Scroll to the channel of the protein whose lifetimes will be assayed.</li>
	<li>Move to frame 10 or 15 of the movie to reduce the inclusion of pre-existing CCPs whose entire durations are not visible. Draw an ROI box around a spot chosen at random.</li>
	<li>Count the number of frames a spot is visible. 
	NOTE: See Figure 3B and 3C for examples of three morphological categories of endocytic events10,11,16-19.</li>
</ol>
4. Analysis of Endocytic Dynamics by Objective Recognition
Objective recognition allows detection of virtually all of the imaged CCPs in a cell, but can be prone to error due to spurious detection of erroneous structures. See discussion for details weighing the advantages and disadvantages of each method. Several programs, including custom-built algorithms, may be used to objectively detect CCPs. This protocol describes objective recognition of CCPs using Imaris, an image-analysis software (see Materials).
<ol>
	<li>To begin, open the raw image file in the image-analysis software. By default, a merged color image appears. Use the &#8220;Display Adjustment&#8221; window to adjust the brightness of a channel. Click on the name of a channel to change the displayed color. Uncheck the box next to the channel to prevent its display (See Figure 4A).</li>
	<li>Create &#8220;Spots&#8221; by clicking on the icon with orange spots. See Figure 4B. Select a Region of Interest (ROI) around the cell. See Figure 4C. Use a ROI to exclude areas that will incorrectly detect bright leading edges and plaques. Analyze multiple cells with different expression levels separately.</li>
	<li>Measure the diameter of a Spot to provide initial estimates for the algorithm. Switch to &#8220;Slice&#8221; mode, and click on polar ends of a spot to measure its diameter. See Figure 4E. Do this for 4 or 5 round Spots that look indicative of a CCP at the height of its stability, after formation and before scission. Use these measurements to calculate average diameter down to the nearest tenth of a micron.</li>
	<li>Detect Spots. Return to &#8220;Surpass&#8221; mode. Select the appropriate channel for detection. Enter the average measured diameter, usually 0.3 or 0.4 &#956;m. Click the blue forward arrow to quantify the Spots. See Figure 4E. If the diameter of Spots is underestimated, spurious points of fluorescence will be identified as Spots. If the diameter is too large, true Spots will be ignored. When unsure, estimate a little bit lower, and filter out the incorrect detections later.</li>
	<li>Filter Spots by Quality. Adjust the filter to capture as many Spots as possible without background. The Quality filter is selected by default20. See Figure 4F.
	<ol>
		<li>Use the &#8220;Quality&#8221; curve as a guide to set an appropriate threshold. Move the bottom threshold of the filter with the left mouse button to this first dip in the curve. This is a good starting point for the filter, as this position usually refines detection to only visible Spots. See Figure 4F.</li>
		<li>Detected Spots appear on the movie as spheres or center pixels. Using the &#8220;Colors&#8221; tab, adjust the color of the Spots to a contrasting color to make it easier to see them. Scroll through the movie to review the Spots in each frame. The goal is to detect as many Spots as possible for the entirety of their lifetimes.</li>
		<li>Eliminate dimmer Spots or manually connect them into continuous tracks later on, as they are not detected well through the course of their lifetimes.</li>
		<li>Remove bright plaques with an &#8220;Intensity&#8221; filter, if required. Click the green plus sign to add a new filter. See Figure 4F. Create an Intensity filter to filter in or out Spots of a given fluorescence signals. Use the generated Intensity curve, (analogous to Quality Curve discussed in 4.5) to set the threshold. Use the left mouse button to remove the extreme left end of the curve that represents the brightest Spots. 
		NOTE: Large plaque structures are usually among the brightest elements in the cell, and they are easily excluded with this filter.</li>
		<li>Scroll through the movie to ensure that the CCPs are detected as Spots and the brightest plaques are no longer detected. Figure 3D shows a cell where detection of CCPs (denoted by white spheres) was optimized with the quality filter and plaques were excluded with the quality filter.</li>
		<li>Reduce bright cell edges with the quality filter. Bright objects may still need to be removed manually, at the next step. Continue with the blue arrow.</li>
		<li>Remove aberrant Spots in the Edit Spots screen. Switch to &#8220;Select&#8221; mode. Click on the spot. It will turn yellow. Click &#8220;Delete&#8221;. Click the blue arrow to continue building the algorithm. See Figure 4G.</li>
	</ol>
	</li>
	<li>Measure tracks. Set tracks to &#8220;Brownian motion.&#8221; The CCP should not exhibit any directed movement, only minimal drifting across the plasma membrane. This algorithm is most appropriate for that motion. See Figure 4H.
	<ol>
		<li>Set the Max Distance to a small value such as 0.7 &#181;m. See Figure 4H. 
		NOTE: Setting a small distance minimizes connection of adjacent Spots and still allows for continued tracking of a cell spot through CCP or cell movement.</li>
		<li>Set Max Gap Size to 1. This allows a track to continue if there is only one frame missing in succession. Click the blue arrow to calculate tracks. See Figure 4H. 
		NOTE: Gap sizes of more than 3 causes different tracks to be incorrectly linked together.</li>
		<li>Switch track viewing to &#8220;Dragon Tail.&#8221; Scroll through the movie observing the detection quality. If adjacent Spots are being connected, decrease the Max Distance below 0.7 &#181;m. If Spots are being broken up too readily, increase the Max Gap Size. Click the blue arrow to create tracks. This leads to the filter tracks screen. See Figure 4H.</li>
	</ol>
	</li>
	<li>Filter Tracks and Export Data. Use an &#8220;Intensity&#8221; filter to remove additional bright plaques. Use a &#8220;Displacement&#8221; filter to remove endosomes that enter the TIRF field. Use caution, as longer Spots will have longer displacements as well. Click the green double arrow to complete the protocol. See Figure 4I.
	<ol>
		<li>Use the Edit Tracks tab, with the pencil icon, to remove false positive detections of Tracks that could not be removed by filters. Check for discontinuities or false connections. To connect two tracks, select them using Ctrl-Left-Click, then click on &#8220;Connect&#8221; in the Edit Tracks box. To disconnect tracks into separate spots, select a track and hit &#8220;Disconnect&#8221;. Select multiple Spots with Ctrl+Left-Click and click &#8220;Connect&#8221; to create a new Track. The method is the same for Editing Spots, described in 4.5.7. See Figure 4J.</li>
		<li>To export data, go to the statistics tab. Click on the single floppy disk icon to export the selected statistic. Click on the icon that looks like a series of floppy disks to export all statistics. The &#8220;Track Duration&#8221; statistic contains the length of the endocytic tracks. See Figure 4K.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here we describe the use of TIRFM to visualize clathrin-mediated endocytosis (CME) at the level of individual CCPs in living cells in real time. CME is a rapid and highly dynamic event mediated by the cumulative effect of many spatially and temporally separate individual events. Most assays that are currently used, such as biochemical measurements of internalization using surface biotinylation or ligand binding, flow-cytometry or fixed cells assays measuring amount of internalized proteins, or electron microscopic locali...

The process of clathrin-mediated endocytosis (CME) is dependent upon the well-timed arrival of the many components of the clathrin-mediated endocytic machinery to gather cargo and manipulate the plasma membrane into the vesicles1-3. CME is initiated by membrane deforming and cargo-adaptor proteins that come together at nascent sites of endocytosis1. These proteins recruit the coat protein clathrin, which assembles into a cage-like structure that forms the clathrin-coated pit (CCP)4. Once the CCP is fully assembled into a spherical shape, membrane scission, primarily through action of the large GTPase, dynamin, generates free clathrin-coated vesicles (CCVs)5,6. This internalization triggers rapid disassembly of the clathrin coat, allowing components to be re-used for multiple rounds of CME.
The discovery and characterization of the proteins involved in CME has been rooted in traditional biochemical, genetic, and microscopy techniques4-6,8. These assays have elucidated the roles and interaction points of these endocytic components. Although very useful for defining essential components of trafficking machinery, these assays are highly limited in capturing the dynamic behavior of CME components or cargo concentration. This is a critical limitation, since CME is driven by the choreographed assembly of sets of protein modules in defined steps, and since small changes in the dynamics of individual endocytic events can have large cumulative consequences on endocytosis. Further, recent data indicate that individual CCPs might differ both in composition and in behavior, suggesting that the physiological regulation of this process is highly spatially and temporally constrained9-14. Visualizing individual endocytic events, therefore, is essential to understand why there are multiple redundant proteins involved in CME and how these proteins might be controlled by physiological signals to regulate cargo internalization.
Here we describe the use of Total Internal Reflection Fluorescence Microscopy (TIRFM) to observe CME at the level of the dynamics of individual CCPs in living cells. TIRFM relies on the difference in refractive index between the glass coverslip and the fluid environment of cells15,16. When the excitation light is directed towards the cells at more than the critical angle, it is internally reflected, creating an evanescent wave that maintains a thin field of illumination extending approximately 100 nm above the coverslip. This ensures that only the fluorescent molecules within this narrow field are excited. Practically, this allows the excitation of fluorescent molecules on or near the plasma membrane, and minimizes fluorescence from the interior parts of the cell. This provides a significantly higher signal-to-noise ratio and z-axis resolution to visualize events at the plasma membrane, compared to more commonly used modes such as conventional epifluorescence or confocal fluorescence microscopy. We also describe, at an introductory and practical level, the use of a commonly used image analysis platform to analyze and quantitate simple morphological features and dynamics of individual cargo endocytic events.

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			<td style="width:157px;">Catalog Number</td>
			<td style="width:207px;">Comments/Description</td>
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			<td height="40" style="height:40px;width:367px;">DMEM/High Glucose with L-glutamine and sodium pyruvate</td>
			<td style="width:180px;">Fisher Scientific&#160;</td>
			<td>SH3024301</td>
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			<td height="40" style="height:40px;width:367px;">Dulbecco&#39;s Phosphate Buffered Saline (DPBS), no calcium, no magnesium</td>
			<td style="width:180px;">Gibco, by Life Technologies</td>
			<td>21600-010</td>
			<td></td>
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			<td height="40" style="height:40px;width:367px;">EDTA Free Acid</td>
			<td style="width:180px;">Amresco</td>
			<td style="width:157px;">0322-500G</td>
			<td></td>
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			<td height="40" style="height:40px;width:367px;">Fetal Bovine Serum</td>
			<td style="width:180px;">Gibco, by Life Technologies</td>
			<td style="width:157px;">10437-028</td>
			<td></td>
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			<td height="40" style="height:40px;width:367px;">Leibovitz&#39;s L-15 Medium, no phenol red</td>
			<td style="width:180px;">Gibco, by Life Technologies</td>
			<td>21083-027</td>
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			<td height="40" style="height:40px;width:367px;">Opti-MEM</td>
			<td style="width:180px;">Gibco, by Life Technologies</td>
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			<td height="40" style="height:40px;width:367px;">HEPES</td>
			<td style="width:180px;">CellPURE by Fisher Scientific&#160;</td>
			<td>BP2937-100</td>
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			<td height="40" style="height:40px;width:367px;">Effectene&#160;</td>
			<td style="width:180px;">Qiagen</td>
			<td style="width:157px;">301425</td>
			<td>Transfection reagent</td>
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			<td height="40" style="height:40px;width:367px;">25 millimeter coverglass</td>
			<td style="width:180px;">Fisher Scientific&#160;</td>
			<td>12-545-86&#160;</td>
			<td></td>
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			<td height="40" style="height:40px;width:367px;">Corning cell culture treated flasks, 25cm2</td>
			<td style="width:180px;">Fisher Scientific&#160;</td>
			<td>10-126-28</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;width:367px;">Cell culture 6-well plate</td>
			<td style="width:180px;">Greiner Bio-One, by VWR</td>
			<td>82050-896</td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;width:367px;">Monoclonal ANTI-FLAG M1</td>
			<td style="width:180px;">Sigma Aldrich</td>
			<td>F3040-5MG</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:367px;">[D-Ala2, N-Me-Phe4, Gly5-ol]-Enkephalin acetate salt (DAMGO)</td>
			<td style="width:180px;">Sigma Aldrich</td>
			<td>E7384-5MG</td>
			<td></td>
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			<td height="40" style="height:40px;width:367px;">Alexa Fluor 647 Protein Labeling Kit</td>
			<td style="width:180px;">Life Technologies</td>
			<td style="width:157px;">A20173</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:367px;">ImageJ</td>
			<td style="width:180px;">NIH</td>
			<td style="width:157px;"></td>
			<td>http://rsb.info.nih.gov/ij/</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:367px;">Imaris Image analysis software</td>
			<td style="width:180px;">BitPLane</td>
			<td style="width:157px;"></td>
			<td>http://www.bitplane.com/imaris/imaris, for automated analysis</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:367px;">Nikon Eclipse Ti inverted microscope and required accessories including filter cubes and filters</td>
			<td style="width:180px;">Nikon</td>
			<td style="width:157px;"></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:367px;">Nikon TIRF arm with required adapters for Nikon Eclipse Ti</td>
			<td style="width:180px;">Nikon</td>
			<td style="width:157px;"></td>
			<td>For adjusting angle of incidence</td>
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			<td height="40" style="height:40px;width:367px;">iXon+ EMCCD camera and adapters</td>
			<td style="width:180px;">Andor</td>
			<td style="width:157px;"></td>
			<td></td>
		</tr>
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visualizing clathrin-mediated endocytosis of g protein-coupled receptors at single-event resolution via tirf microscopy

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological assays, measuring global changes in endocytosis, have identified over 30 known components participating in CME, and biochemical studies have generated an interaction map of many of these components. It is becoming increasingly clear, however, that CME is a highly dynamic process whose regulation is complex and delicate. In this manuscript, we describe the use of Total Internal Reflection Fluorescence (TIRF) microscopy to directly visualize the dynamics of components of the clathrin-mediated endocytic machinery, in real time in living cells, at the level of individual events that mediate this process. This approach is essential to elucidate the subtle changes that can alter endocytosis without globally blocking it, as is seen with physiological regulation. We will focus on using this technique to analyze an area of emerging interest, the role of cargo composition in modulating the dynamics of distinct clathrin-coated pits (CCPs). This protocol is compatible with a variety of widely available fluorescence probes, and may be applied to visualizing the dynamics of many cargo molecules that are internalized from the cell surface.

Using live-cell TIRF Microscopy we have recorded the endocytic dynamics of the &#181;-opioid-receptor (MOR), a G-protein coupled receptor (GPCR) and its endocytic adaptor protein &#946;-arrestin. The &#946;-arrestin construct was transiently transfected into a stably expressing MOR cell line using the protocol outlined in Figure 1, and imaged 96 hr later. The MOR in the stable cell line is N-terminally tagged with a pH-sensitive GFP. This fluorescent protein only fluoresces in the neutral extracellular f...

Watch this Scientific Journal Video about Visualizing Clathrin-mediated Endocytosis of G Protein-coupled Receptors at Single-event Resolution via TIRF Microscopy at JoVE.com

Visualizing Clathrin-mediated Endocytosis of G Protein-coupled Receptors at Single-event Resolution via TIRF Microscopy

Clathrin-mediated endocytosis, a rapid and highly dynamic process internalizes many proteins, including signaling receptors. The protocol described here directly visualizes the kinetics of individual endocytic events. This is essential for understanding how core members of the endocytic machinery coordinate with each other, and how protein cargo influence this process.

Many important signaling receptors are internalized through the well-studied process of clathrin-mediated endocytosis (CME). Traditional cell biological ...

visualizing-clathrin-mediated-endocytosis-g-protein-coupled-receptors

Research

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Biology

80.1K 조회수.  Carnegie Mellon University. Clathrin-mediated endocytosis, a rapid and highly dynamic process internalizes many proteins, including signaling receptors. The protocol described here directly visualizes the kinetics of individual endocytic events. This is essential for understanding how core members of the endocytic machinery coordinate with each other, and how protein cargo influence this process. 

비디오: Visualizing Clathrin-mediated Endocytosis of G Protein-coupled Receptors at Single-event Resolution via TIRF Microscopy

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. F&ouml;rster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 &alpha;-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

By employing a spectrally resolved two-photon microscopy imaging system, pixel-level maps of F&ouml;rster Resonance Energy Transfer (FRET) efficiencies are obtained for cells expressing membrane receptors hypothesized to form homo-oligomeric complexes. From the FRET efficiency maps, we are able to estimate stoichiometric information about the oligomer complex under study.

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Imaging G-protein Coupled Receptor (GPCR)-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

Many eukaryotic cells can detect gradients of chemical signals in their environments and migrate accordingly 1. This guided cell migration is referred as chemotaxis, which is essential for various cells to carry out their functions such as trafficking of immune cells and patterning of neuronal cells 2, 3. A large family of G-protein coupled receptors (GPCRs) detects variable small peptides, known as chemokines, to direct cell migration in vivo 4. The final goal of chemotaxis research is to understand how a GPCR machinery senses chemokine gradients and controls signaling events leading to chemotaxis. To this end, we use imaging techniques to monitor, in real time, spatiotemporal concentrations of chemoattractants, cell movement in a gradient of chemoattractant, GPCR mediated activation of heterotrimeric G-protein, and intracellular signaling events involved in chemotaxis of eukaryotic cells 5-8. The simple eukaryotic organism, Dictyostelium discoideum, displays chemotaxic behaviors that are similar to those of leukocytes, and D. discoideum is a key model system for studying eukaryotic chemotaxis. As free-living amoebae, D. discoideum cells divide in rich medium. Upon starvation, cells enter a developmental program in which they aggregate through cAMP-mediated chemotaxis to form multicullular structures. Many components involved in chemotaxis to cAMP have been identified in D. discoideum. The binding of cAMP to a GPCR (cAR1) induces dissociation of heterotrimeric G-proteins into G&gamma; and G&beta;&gamma; subunits 7, 9, 10. G&beta;&gamma; subunits activate Ras, which in turn activates PI3K, converting PIP2 into PIP3 on the cell membrane 11-13. PIP3 serve as binding sites for proteins with pleckstrin Homology (PH) domains, thus recruiting these proteins to the membrane 14, 15. Activation of cAR1 receptors also controls the membrane associations of PTEN, which dephosphorylates PIP3 to PIP2 16, 17. The molecular mechanisms are evolutionarily conserved in chemokine GPCR-mediated chemotaxis of human cells such as neutrophils 18. We present following methods for studying chemotaxis of D. discoideum cells. 1. Preparation of chemotactic component cells. 2. Imaging chemotaxis of cells in a cAMP gradient. 3. Monitoring a GPCR induced activation of heterotrimeric G-protein in single live cells. 4. Imaging chemoattractant-triggered dynamic PIP3 responses in single live cells in real time. Our developed imaging methods can be applied to study chemotaxis of human leukocytes.

Imaging G-protein Coupled Receptor GPCR-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

Here, we describe detailed live cell imaging methods for investigating chemotaxis. We present fluorescence microscopic methods to monitor spatiotemporal dynamics of signaling events in migrating cells. Measurement of signaling events permits us to further understand how a GPCR-signaling network achieves gradient sensing of chemoattractants and controls directional migration of eukaryotic cells.

Many eukaryotic cells can detect gradients of chemical signals in their environments and migrate accordingly 1.  This guided cell migration is referred as ...

Quantifying Agonist Activity at G Protein-coupled Receptors

When an agonist activates a population of G protein-coupled receptors (GPCRs), it elicits a signaling pathway that culminates in the response of the cell or tissue. This process can be analyzed at the level of a single receptor, a population of receptors, or a downstream response. Here we describe how to analyze the downstream response to obtain an estimate of the agonist affinity constant for the active state of single receptors.
Receptors behave as quantal switches that alternate between active and inactive states (Figure 1). The active state interacts with specific G proteins or other signaling partners. In the absence of ligands, the inactive state predominates. The binding of agonist increases the probability that the receptor will switch into the active state because its affinity constant for the active state (Kb) is much greater than that for the inactive state (Ka). The summation of the random outputs of all of the receptors in the population yields a constant level of receptor activation in time. The reciprocal of the concentration of agonist eliciting half-maximal receptor activation is equivalent to the observed affinity constant (Kobs), and the fraction of agonist-receptor complexes in the active state is defined as efficacy (&#949;) (Figure 2).
Methods for analyzing the downstream responses of GPCRs have been developed that enable the estimation of the Kobs and relative efficacy of an agonist 1,2. In this report, we show how to modify this analysis to estimate the agonist Kb value relative to that of another agonist. For assays that exhibit constitutive activity, we show how to estimate Kb in absolute units of M-1.
Our method of analyzing agonist concentration-response curves 3,4 consists of global nonlinear regression using the operational model 5. We describe a procedure using the software application, Prism (GraphPad Software, Inc., San Diego, CA). The analysis yields an estimate of the product of Kobs and a parameter proportional to efficacy (&tau;). The estimate of &tau;Kobs of one agonist, divided by that of another, is a relative measure of Kb (RAi) 6. For any receptor exhibiting constitutive activity, it is possible to estimate a parameter proportional to the efficacy of the free receptor complex (&tau;sys). In this case, the Kb value of an agonist is equivalent to &tau;Kobs/&tau;sys 3.
Our method is useful for determining the selectivity of an agonist for receptor subtypes and for quantifying agonist-receptor signaling through different G proteins.

A method for estimating the affinity constant of an agonist for the active state (Kb) of a G protein-coupled receptor is described. The analysis provides absolute or relative measures of Kb depending on whether constitutive receptor activation is measurable. Our method applies to various responses downstream from receptor activation.

When an agonist activates a population of G protein-coupled receptors (GPCRs), it elicits a signaling pathway that culminates in the response of the cell ...

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Genetically-encoded Molecular Probes to Study G Protein-coupled Receptors

To facilitate structural and dynamic studies of G protein-coupled receptor (GPCR) signaling complexes, new approaches are required to introduce informative probes or labels into expressed receptors that do not perturb receptor function. We used amber codon suppression technology to genetically-encode the unnatural amino acid, p-azido-L-phenylalanine (azF) at various targeted positions in GPCRs heterologously expressed in mammalian cells. The versatility of the azido group is illustrated here in different applications to study GPCRs in their native cellular environment or under detergent solubilized conditions. First, we demonstrate a cell-based targeted photocrosslinking technology to identify the residues in the ligand-binding pocket of GPCR where a tritium-labeled small-molecule ligand is crosslinked to a genetically-encoded azido amino acid. We then demonstrate site-specific modification of GPCRs by the bioorthogonal Staudinger-Bertozzi ligation reaction that targets the azido group using phosphine derivatives. We discuss a general strategy for targeted peptide-epitope tagging of expressed membrane proteins in-culture and its detection using a whole-cell-based ELISA approach. Finally, we show that azF-GPCRs can be selectively tagged with fluorescent probes. The methodologies discussed are general, in that they can in principle be applied to any amino acid position in any expressed GPCR to interrogate active signaling complexes.

We genetically-encode the unnatural amino acid, p-azido-L-phenylalanine at various targeted positions in GPCRs and show the versatility of the azido group in different applications. These include a targeted photocrosslinking technology to identify residues in the ligand-binding pocket of a GPCR, and site-specific bioorthogonal modification of GPCRs with a peptide-epitope tag or fluorescent probe.

To facilitate structural and dynamic studies of G protein-coupled receptor (GPCR) signaling complexes, new approaches are required to introduce ...

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

Characterization of G Protein-coupled Receptors by a Fluorescence-based Calcium Mobilization Assay

For more than 20 years, reverse pharmacology has been the preeminent strategy to discover the activating ligands of orphan G protein-coupled receptors (GPCRs). The onset of a reverse pharmacology assay is the cloning and subsequent transfection of a GPCR of interest in a cellular expression system. The heterologous expressed receptor is then challenged with a compound library of candidate ligands to identify the receptor-activating ligand(s). Receptor activation can be assessed by measuring changes in concentration of second messenger reporter molecules, like calcium or cAMP. The fluorescence-based calcium mobilization assay described here is a frequently used medium-throughput reverse pharmacology assay. The orphan GPCR is transiently expressed in human embryonic kidney 293T (HEK293T) cells and a promiscuous G&#945;16 construct is co-transfected. Following ligand binding, activation of the G&#945;16 subunit induces the release of calcium from the endoplasmic reticulum. Prior to ligand screening, the receptor-expressing cells are loaded with a fluorescent calcium indicator, Fluo-4 acetoxymethyl. The fluorescent signal of Fluo-4 is negligible in cells under resting conditions, but can be amplified more than a 100-fold upon the interaction with calcium ions that are released after receptor activation. The described technique does not require the time-consuming establishment of stably transfected cell lines in which the transfected genetic material is integrated into the host cell genome. Instead, a transient transfection, generating temporary expression of the target gene, is sufficient to perform the screening assay. The setup allows medium-throughput screening of hundreds of compounds. Co-transfection of the promiscuous G&#945;16, which couples to most GPCRs, allows the intracellular signaling pathway to be redirected towards the release of calcium, regardless of the native signaling pathway in endogenous settings. The HEK293T cells are easy to handle and have proven their efficacy throughout the years in receptor deorphanization assays. However, optimization of the assay for specific receptors may remain necessary.

The here described fluorescence-based calcium mobilization assay is a medium-throughput reverse pharmacology screening system for the identification of functionally activating ligand(s) of orphan G protein-coupled receptors (GPCRs).

For more than 20 years, reverse pharmacology has been the preeminent strategy to discover the activating ligands of orphan G protein-coupled receptors ...