Our work is supported by the Biotechnology and Biological Sciences Research Council. We would like to thank Prof Tamotsu Yoshimori for kindly supplying us with the plasmid for the expression of CFP-LC3.

1. Cell Preparation
<ol>
 <li>Seed low passage number of HEK-293T cells stably expressing GFP-DFCP1 on 22 mm round coverslips; culture cells overnight in Dulbecco Modified Eagles's Medium (DMEM), to a confluency of 30-40% (aim for a confluency of 80% after 2 days - day of live cell imaging).</li>
</ol>
2. Cell Transfection
<ol>
 <li>Prepare the transfection complex mix for each plate, containing 100 &mu;l OptiMEM I reduced serum medium, 3 &mu;l X-tremeGENE 9 DNA Transfection Reagent, and 0.5 &mu;g of pECFP-LC3 plasmid DNA. Mix gently by pipetting up and down and incubate 15 min at room temperature. (Note: we have repeatedly found that other transfection reagents, such as Lipofectamine 2000, have a lot of toxicity, and, in addition, they produce fluorescent particles on their own which interfere with many microscopy techniques.)</li>
 <li>Aspirate the medium from plates and add fresh DMEM pre-warmed at 37 &deg;C.</li>
 <li>Add the transfection complex to cells by pipetting; incubate cells for 24 hr.</li>
</ol>
3. Cell Incubation with the Organelle Marker (Optional)
<ol>
 <li>Add mitotracker/lysotracker to DMEM at a final concentration of 75 nM, and keep on ice, in a falcon tube covered with aluminum foil, along the whole experimental day, to avoid light exposure and repetitive freezing and thawing cycles.</li>
 <li>Remove an aliquot of 2 ml of the mitotracker/lysotracker-containing medium and warm up at 37 &deg;C; aspirate medium from transfected cells and replace with mitotracker/ lysotracker-containing medium and incubate cells for 30 - 60 min.</li>
</ol>
4. Preparation of Incubation Chamber for Live Cell Imaging (Figure 1)
<ol>
 <li>Clean metal and plastic O-rings with 75% ethanol and apply silicon grease on the rim of the metal O-ring.</li>
 <li>Using forceps remove the coverslip from the plate and dry the excess of medium from the bottom side of the coverslip, to avoid mixing too much medium with the grease, as this will increase the likelihood of leakage.</li>
 <li>Leave the coverslip to rest on the ledge of the plastic O-ring and fit the metal O-ring on top of the plastic O-ring, with the coverslip sandwiched in between, in order to create a closed chamber.</li>
 <li>Top up chamber with the medium from the plate; from this point and on, avoid prolonged incubation of the cells in DMEM medium without a buffering agent, to prevent changes in pH to occur, as the bicarbonate buffer system of DMEM requires artificial CO2 concentration of 5-10% and the CO2 concentration of ambient air is much lower.</li>
</ol>
5. Starvation of Cells
<ol>
 <li>Put the incubation chamber on the microscope stage.</li>
 <li>Aspirate the complete medium and wash with 2 ml of starvation medium 3 times, to ensure that no amino acids have remained from the DMEM, which will eventually inhibit the autophagy response; set the timer ON.</li>
</ol>
6. Microscopy
<ol>
 <li>An appropriate imaging system configured for live cell wide-field epi-fluorescence will be required. This will typically comprise a research-grade inverted microscope frame, an intense broad-spectrum light source, mirrors and filters specific for the fluorescent protein(s)/dye(s) of interest, a high quality objective lens, a sensitive CCD/sCMOS camera and an incubation chamber. All the major microscope manufacturers offer complete wide-field systems appropriate for live cell imaging, but it is also possible to home-build a system using components from a variety of manufacturers and control it using open source software such as Micro-Manager (<a href="http://valelab.ucsf.edu/~MM/MMwiki/" target="_blank">http://valelab.ucsf.edu/~MM/MMwiki/</a>). The most important aspect in our opinion is to use a system of high sensitivity so that expression levels of the fluorescent reporters can be kept to a minimum.</li>
 <li>Selecting the appropriate cells to image.
 <ol>
 <li>Select large and flat cells that will allow more autophagosome formation events to be captured. Also, opt for cells that have already started producing a greater number of omegasomes.</li>
 <li>Start the video capture after 30 min or well into autophagy response, in order to capture a larger ratio of autophagosome formation events per video.</li>
 </ol>
 </li>
 <li>Imaging.
 <ol>
 <li>Use a high magnification lens (100x 1.4 NA).</li>
 <li>Adjust the intensity of excitation light to 10-20% of maximum to prevent photo-bleaching.</li>
 <li>Set the camera (Hamamatsu ORCA ER, pixel size 6.45 &mu;m) to 100-500 msec exposure, 2x2 binning and 100 gain.</li>
 <li>Set the image acquisition rate to 1 frame every 10 sec.</li>
 </ol>
 </li>
</ol>
7. Creating Montages of Autophagosome Formation Events with ImageJ
(Note: This can be done in a non-systematic way by simply scanning the merged video for events of interest, but it can also be systematized as outlined below.)
<ol>
 <li>Open image stacks for the 3 (or 2) captured channels in ImageJ/Fiji.</li>
 <li>Apply green, red and blue LUT (Lookup Tables) to the corresponding channel; merge 3 colors and save.</li>
 <li>From the Analyze tab, select Tools &gt; ROI manager... &gt; Specify, then select a random area of defined size in the first image of the stack.</li>
 <li>From the Image tab, select duplicate, in order to duplicate the selected area of the stack.</li>
 <li>From the Image tab, select Stacks &gt; Make montage... to create a tentative montage containing all the frames captured. Scan all frames in the montage for a complete autophagosome formation event; keep notes of the first and last frame of the event.</li>
 <li>From the Analyze tab, select Tools &gt; ROI manager... select the same area in the first image of the stack for the other 2 colors as well as for the merged colors image and duplicate stacks.</li>
 <li>From the File tab, select New &gt; Image, and set for width according to the number of pixels initially selected using the ROI manager, set for height 4 times the height set in the ROI manager plus 3 pixels for space in between the 3 colors and the merged, and set Slices as the number of frames in each stack.</li>
 <li>From the Image tab, select Stacks &gt; Tools &gt; Insert..., then insert each sub-stack one on top of another, leaving space of 1 pixel in between.</li>
 <li>From the Image tab, select Stacks &gt; Make montage... to create a montage starting and finishing with the first and last frame of the autophagosome formation event captured.</li>
</ol>

<ol>
	<li>Mizushima, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autophagy:+process+and+function.">Autophagy: process and function.</a> Genes Dev. 21, 2861-2873 (2007).</li><li>Mizushima, N., Yoshimori, T., Ohsumi, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+Atg+proteins+in+autophagosome+formation.">The role of Atg proteins in autophagosome formation.</a> Annual review of cell and developmental biology. 27, 107-132 (2011).</li><li>Klionsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autophagy:+from+phenomenology+to+molecular+understanding+in+less+than+a+decade.">Autophagy: from phenomenology to molecular understanding in less than a decade.</a> Nat. Rev. Mol. Cell Biol. 8, 931-937 (2007).</li><li>Klionsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autophagy+revisited:+a+conversation+with+Christian+de+Duve.">Autophagy revisited: a conversation with Christian de Duve.</a> Autophagy. 4, 740-743 (2008).</li><li>Yla-Anttilba, P., Vihinen, H., Jokitalo, E., Eskelinen, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+tomography+reveals+connections+between+the+phagophore+and+endoplasmic+reticulum.">3D tomography reveals connections between the phagophore and endoplasmic reticulum.</a> Autophagy. 5, 1180-1185 (2009).</li><li>Hayashi-Nishino, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+subdomain+of+the+endoplasmic+reticulum+forms+a+cradle+for+autophagosome+formation.">A subdomain of the endoplasmic reticulum forms a cradle for autophagosome formation.</a> Nat. Cell Biol. 11, 1433-1437 (2009).</li><li>Lippincott-Schwartz, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emerging+in+vivo+analyses+of+cell+function+using+fluorescence+imaging+(*).">Emerging in vivo analyses of cell function using fluorescence imaging (*).</a> Annu. Rev. Biochem. 80, 327-332 (2011).</li><li>Mizushima, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissection+of+autophagosome+formation+using+Apg5-deficient+mouse+embryonic+stem+cells.">Dissection of autophagosome formation using Apg5-deficient mouse embryonic stem cells.</a> The Journal of Cell Biology. 152, 657-668 (2001).</li><li>Itakura, E., Mizushima, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+autophagosome+formation+site+by+a+hierarchical+analysis+of+mammalian+Atg+proteins.">Characterization of autophagosome formation site by a hierarchical analysis of mammalian Atg proteins.</a> Autophagy. 6, 764-776 (2010).</li><li>Axe, E. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autophagosome+formation+from+membrane+compartments+enriched+in+phosphatidylinositol+3-phosphate+and+dynamically+connected+to+the+endoplasmic+reticulum.">Autophagosome formation from membrane compartments enriched in phosphatidylinositol 3-phosphate and dynamically connected to the endoplasmic reticulum.</a> J Cell Biol. 182, 685-701 (2008).</li><li>Walker, S., Chandra, P., Manifava, M., Axe, E., Ktistakis, N. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Making+autophagosomes:+localized+synthesis+of+phosphatidylinositol+3-phosphate+holds+the+clue.">Making autophagosomes: localized synthesis of phosphatidylinositol 3-phosphate holds the clue.</a> Autophagy. 4, 1093-1096 (2008).</li></ol>

No conflicts of interest declared.

The method described in this protocol allows the visualization of a protein's localization during autophagosome formation. We have tried various methods of visualizing the events described including point-scanning confocal, spinning disk confocal and Total Internal Reflection Fluorescence (TIRF) microscopy. We have found that for general purposes standard wide-field epi-fluorescence provides the best compromise between sensitivity and resolution. This ensures good signal to noise, minimal photo-bleaching/photo-toxicity and fast acquisition. The lack of optical sectioning is not an issue if appropriate regions of the cell are chosen to image i.e. the periphery where the cell is spread and flat. However, it is important that the imaging system used is appropriately configured (both in terms of the hardware used and the system settings).	
	For best spatial resolution, it is recommended to use a high magnification, high numerical aperture oil immersion lens (water immersion lenses will offer no benefit imaging at close proximity to the coverslip). It is suggested to balance the intensity of illumination (e.g. with neutral density filters), the camera settings (exposure time, binning and gain) to maximize the signal-to-noise and minimize bleaching. This will have to be done empirically, but as a guide, when using a 100x 1.4 NA lens we typically reduce the power of our excitation light to 10-20% of maximum and set the camera (Hamamatsu ORCA ER, pixel size 6.45 &mu;m) to 100-500 msec exposure, 2x2 binning and 100 gain.	
	The image acquisition rate should be set in the range of 1 frame every 1-10 sec. Acquiring images at higher frame rates will ensure better continuity between images (better temporal resolution) but will expose cells to more light and thus increase photo-bleaching/photo-toxicity.	
	If imaging more fluorescence channels, it has to be ensured that the delay between channel capture is minimized (reduce exposure time, fit fast filter changers). This will reduce the chances of motion artifacts appearing in the composite image. If motion artifacts are proving difficult to avoid consider using an image splitter (a device to facilitate the simultaneous acquisition of two fluorescence channels using one camera) or a dual camera adapter.	
	Imaging blue channel requires the selection of suitable filters and mirrors to prevent cross-emission of the blue fluorescence to the green channel. We have successfully used an Olympus CellR microscope, which uses the Till Polychrome V illuminator, enabling specific selection of wavelength, bandwidth and intensity and so is very flexible. However, the light source is 'leaky' with some white light coming through in addition to the selected wavelengths.For this reason we have fitted (multi) bandpass excitation filters in the cubes, so there is additional filtering of the excitation light. The following combination of mirrors/filters was used (all Semrock). For GFP &amp; mCherry: Exciter FF01-479-585, Emitter FF02-525/40 (GFP) and FF01-607/36 (mCherry), Dichroic Mirror FF505/606-Di01. For CFP: Exciter FF01-416/501, Emitter FF01-523/610, Dichroic Mirror FF440/520-Di01. We have also used a different CellR microscope, which has given sub-optimal results, with cross-emission of blue fluorescence to the green channel. This microscope uses a white light source and has a fast filter wheel to select the excitation wavelengths.This means that the wavelength selection is limited to the 8 filters in the wheel, but there is a separate wheel to regulate intensity. The following combination of mirrors/filters was used. For GFP (Semrock): Exciter FF01-470/40, Emitter FF02-525/50, Dichroic Mirror FF495-Di02. For CFP and mCherry: Exciter FF01-427/10 (CFP, Semrock) 572/23 (mCherry, Chroma), Emitter FF01-472/30 (CFP, Semrock) 632/60 (mCherry, Chroma), Dichroic Mirror 89006bs (Chroma).	
	The significance of this technique compared to other imaging techniques is two-fold: First, it can capture the localization of the protein of interest in live cells, and second, it can increase the information extracted adding the fourth dimension of time. However, as with exogenous proteins there is always the possibility of mislocalization, either due to increased expression levels or due to tagging, therefore live cell imaging should be combined with immuno-staining of the endogenous POI in fixed cells, in order to corroborate the results. Finally, it is worthwhile noting that live cell imaging can be combined with immuno-EM, in order to increase the spatial resolution of the analysis.

Autophagy is a highly dynamic process, which requires the coordination of a large number of proteins for the final outcome of autophagosome formation1-3. Microscopy is probably the technique most commonly applied for studying autophagy4. The localization of most autophagy proteins has been extensively studied in fixed cells, both by immuno-staining the endogenous proteins and by expression of fluorescently tagged exogenous protein. In addition, Electron Microscopy (EM), alone and in combination with immuno-gold labeling, has described the fine details of these structures5,6 . Despite the fact that these techniques have established our understanding of autophagosome formation in the 3 dimensions of space, they have failed to provide sufficient amount of information about the 4th dimension - time. Live cell imaging overcomes this barrier as it allows following the formation of an autophagosome as close as possible to real time7. This technique was first employed to study autophagy by Yoshimori and colleagues8, and has been increasingly used henceforth.	
	Time-lapse microscopy captures the localization of the POI in live cells and over a period of time. By comparing this information with a well-characterized autophagy and/or organelle marker, live cell imaging analysis can put the POI in the greater spatial and temporal context of autophagosome formation. Live cell imaging analysis is based on the repetitive capturing of the POI localization along all the steps of autophagosome formation, while imaging of fixed cells is based on a single capture. Therefore, live cell imaging can prove the contribution of the POI at specific steps of autophagosome formation, while imaging of fixed cells can only assume the role of POI, based on its average localization in many autophagosomes simultaneously captured at different stages of their lifecycle.	
	Although live cell imaging is a method of high analytical power, it has some inherent limitations, which should be taken into consideration. First of all, live cell imaging requires the expression of one or more exogenous fluorescently labeled proteins. Fluorescent tags tend to be big in size and they can sometimes alter the behavior of a protein due to steric reasons. This situation is accentuated for membrane proteins, as they need to function in the limited space of the 2 dimensions of membranes. Of note, autophagosomes are membranous structures, and accordingly their formation requires a large number of membrane-associated proteins.	
	Another set of problems is connected to the expression levels of the POI. In principle, an exogenous protein should be expressed at levels comparable to the endogenous protein. This ensures that important regulators of its sub-cellular localization will not be saturated, and the analysis will be biologically relevant. Moreover, overexpression of autophagy proteins should be avoided, as when they are expressed above the endogenous levels, they tend to inhibit the autophagy response9. Conversely, since the expression levels of the POI should be high enough to allow following its localization for a good period of time without photo-bleaching, a compromise has to be reached. Achieving the optimal expression levels of an exogenous protein in mammalians cells requires a lot of fine tuning, but it is feasible by establishing and screening cell lines stably expressing different levels of the POI.	
	The spatial resolution that can be achieved with standard fluorescence microscopy is another limiting factor. Resolution may be limited for a number of reasons, but at best, lateral resolution will be around 250 nm. This means that any objects separated by a distance smaller than this will appear connected (or as a single object) and objects smaller than 250 nm will be represented in the image larger than they actually are. Therefore images should always be interpreted with this in mind and complementary techniques such as EM will be required to resolve fine ultra-structural detail.	
	Finally, live cell imaging inherently requires exposing a cell to light, potentially for a prolonged period of time. This may alter the physiological responses of a cell, a phenomenon known as photo-toxicity.	
	We have successfully used live cell imaging of the PI3P-binding protein DFCP1 to describe for the first time that autophagosomes originate from PI3P-rich ring-like structures termed omegasomes, which are in close association with ER strands10,11 . We have clearly shown that LC3-positive structures start forming in close association with omegasomes. We here suggest that employing a cell line stably expressing GFP-DFCP1 for the live cell imaging of the protein of interest, establishes a robust spatial and temporal frame for the characterization of its role in autophagosome formation.

<table border="1">
		<tbody>
		 <tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		 </tr>
		 <tr>
			<td>DMEM</td>
			<td>Invitrogen</td>
			<td>41965</td>
			<td></td>
		 </tr>
		 <tr>
			<td>OptiMEM I</td>
			<td>Invitrogen</td>
			<td>31985-062</td>
			<td></td>
		 </tr>
		 <tr>
			<td>MitoTracker Red FM</td>
			<td>Invitrogen</td>
			<td>M22425</td>
			<td></td>
		 </tr>
		 <tr>
			<td>LysoTracker Red DND-99</td>
			<td>Invitrogen</td>
			<td>L-7528</td>
			<td></td>
		 </tr>
		 <tr>
			<td>X-tremeGENE 9 DNA Transfection Reagent</td>
			<td>Roche Applied Science</td>
			<td>6365787001</td>
			<td></td>
		 </tr>
		 <tr>
			<td>22 mm coverslips</td>
			<td>VWR</td>
			<td>631-0159</td>
			<td></td>
		 </tr>
		 <tr>
			<td>35 mm plates</td>
			<td>Fisher NUNC</td>
			<td>153066</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Silicon grease</td>
			<td>RS Components Ltd. </td>
			<td>RS 494-124</td>
			<td></td>
		 </tr>
		 <tr>
			<td>O-rings</td>
			<td></td>
			<td></td>
			<td>Custom made</td>
		 </tr>
		 <tr>
			<td>Attofluor Cell Chamber</td>
			<td>Invitrogen</td>
			<td>A-7816</td>
			<td>Suggested alternative to custom-made O-rings</td>
		 </tr>
		 <tr>
			<td>Microscope</td>
			<td>Olympus</td>
			<td>IX81</td>
			<td>Inverted microscope</td>
		 </tr>
		 <tr>
			<td>Objective</td>
			<td>Olympus</td>
			<td>UPLSAPO 100XO</td>
			<td>N.A. 1.4, W.D. 0.13, FN 26.5</td>
		 </tr>
		 <tr>
			<td>Camera</td>
			<td>Hamamatsu</td>
			<td>ORCA-R2 C10600 10B</td>
			<td>Progressive scan interline CCD</td>
		 </tr>
		 <tr>
			<td>Illuminator</td>
			<td>TILL Photonics</td>
			<td>Polychrome V</td>
			<td>Ultrafast monochromator</td>
		 </tr>
		 <tr>
			<td>Incubation chamber</td>
			<td>Solent Scientific</td>
			<td>Cell^R IX81</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Software</td>
			<td>Olympus</td>
			<td>SIS xcellence</td>
			<td></td>
		 </tr>
		</tbody>
 </table>

live cell imaging of early autophagy events: omegasomes and beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

In the protocol described, we have used time-lapse microscopy to follow the localization of CFP-tagged LC3 in a cell line stably expressing GFP-tagged DFCP1, under autophagy inducing conditions. The outcome of this experiment is the capture of 2 series or stacks of images, one from the green and one from the blue channel, corresponding to GFP-DFCP1 and CFP-LC3. We have further analyzed these videos using ImageJ, in order to create montages corresponding to single autophagosome formation events, as described in the protocol section. This analysis allowed us to prove that an LC3-positive autophagosome originates from a DFCP1-positive omegasome. In the montage shown in Figure 2, the formation of an omegasome becomes apparent from the second frame, in the form of a small spot. The omegasome starts expanding in order to form the characteristic ring-like structure, and reaches its maximum diameter after 6 min. Next, the omegasome starts collapsing and eventually it disappears, after approximately 10 min. Putting now the formation of the LC3-positive structure or autophagosome in the context of the omegasome formation, we observe that the autophagosome appears after the omegasome, and becomes clearly visible after approximately 1.5 min. The autophagosome starts expanding in close association with the omegasome, first spot and then ring, and when the omegasome starts collapsing the autophagosome buds off. Eventually the autophagosome stays behind, apparently to fuse with the lysosomes, after the omegasome disappears. This analysis provides a clear indication about the functional relationship between the 2 structures, according to which LC3 positive autophagosomes originate from corresponding omegasomes. 
 In another example, we have added a lysosome tracker in order to capture the temporal and spatial association of the forming autophagosome with lysosomes (Figure 3). Indeed, a few minutes after the collapse of the omegasome, the nascent autophagosome starts associating with lysosomes and eventually it fuses with one lysosome. Fusion of the autophagosome with the lysosome largely quenches CFP-LC3 fluorescence, presumably due to the exposure of the CFP tag in the acidic environment of the lysosome. 
 However, the same kind of analysis can produce uninterpretable results, due to a variety of reasons. In the example presented in Figure 4, the results become uninterpretable due to a drift in focus. Video starts successfully capturing the formation of an autophagosome, however a drift in focus occurs after the 3 min mark. The video capture continues out of focus for the next 6 min, but eventually the focus is manually corrected after the 9 to 10 min mark. This analysis though, makes impossible to discriminate whether the autophagosome formation event captured when the focus is corrected is the initial event which is almost completed, or a new one that has started after the drift in focus. 
 Additional problems can be attributed to the confluency of the cultured cells. For instance, when cells are grown at higher than optimal confluency, they are forced to expand on top of an adjacent cell, which makes focusing quite challenging. Moreover, cells that are grown at high confluency tend to be stressed, increasing the levels of background autophagy activity before the initiation of starvation. 
 Finally, fluorescence-related issues are quite common. CFP has weaker fluorescence activity compared to GFP, and often gets photo-bleached at later stages of the video capturing. Analysis of such videos can lead to false negative conclusions about the spatial association of the POI with forming autophagosomes. However, these problems can be overcome by using one of the variants such as mTurquise2. Another common phenomenon stems from the fact that the fluorescence of red tags is not quenched at the lower pH of lysosomes. Many autophagy proteins physiologically finish their lifecycle into lysosomes, while autophagosomes eventually fuse with lysosomes. Moreover, non-functional proteins are often targeted for degradation in lysosomes. Therefore, one might end up following the non-related association of autophagosomes with lysosomes, instead of an actual physical association of between a red-tagged POI and omegasomes. 
 <img src="/files/ftp_upload/50484/50484fig1.jpg" alt="Figure 1"/> 
 Figure 1. Incubation chamber. The plastic O-ring fits around the rim of the metal O-ring and has at the bottom a thin ledge which extends inwards. The coverslip is placed on the ledge of the plastic O-ring, and then the plastic O-ring is fitted at the bottom of the metal O-ring. This way, the coverslip is sandwiched between the 2 O-rings, creating a closed chamber. 
 <img src="/files/ftp_upload/50484/50484fig2.jpg" alt="Figure 2" fo:content-width="6in" fo:src="/files/ftp_upload/50484/50484fig2highres.jpg" /> 
 Figure 2. Cells expressing GFP-DFCP1 and CFP-LC3 were starved for 30 min and imaged at a rate of 1 frame every 10 sec. A montage of a representative autophagosome formation event is presented. Signals from green and blue channels are pseudo-colored green and red correspondingly. Arrows indicate the first discernible omegasome and autophagosome. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50484/50484fig2large.jpg" target="_blank"> Click here to view larger figure</a>. 
 <img src="/files/ftp_upload/50484/50484fig3.jpg" alt="Figure 3" fo:content-width="6in" fo:src="/files/ftp_upload/50484/50484fig3highres.jpg"/> 
 Figure 3. Cells expressing GFP-DFCP1 and CFP-LC3, were incubated with lysotracker red, starved for 30 min and imaged at a rate of 1 frame every 15 sec. A montage of a representative autophagosome formation event is presented. Signals from green, red and blue channels are pseudo-colored green, blue and red correspondingly. Arrows indicate the first discernible omegasome and autophagosome. Arrowhead indicates the first capture of autophagosome fusion with the lysosome. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50484/50484fig3large.jpg" target="_blank"> Click here to view larger figure</a>. 
 <img src="/files/ftp_upload/50484/50484fig4.jpg" alt="Figure 4" fo:content-width="6in" fo:src="/files/ftp_upload/50484/50484fig4highres.jpg"/> 
 Figure 4. Cells expressing GFP-DFCP1 and CFP-LC3 were starved for 30 min and imaged at a rate of 1 frame every 10 sec. A montage of an example of sub-optimal capture is presented. Signals from green and blue channels are pseudo-colored green and red correspondingly. Arrows indicate the first discernible omegasome and autophagosome. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50484/50484fig4large.jpg" target="_blank"> Click here to view larger figure</a>. 
 Video 1. Video of the autophagosome formation event presented in Figure 2. The playback rate is 4 frames per second. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50484/50484movie1.avi" target="_blank"> Click here to view movie</a>. 
 Video 2. Video of the autophagosome formation event presented in Figure 3. The arrow indicates first, the formation of the omegasome, and second, the fusion of the autophagosome with the lysosome. The playback rate is 4 frames per second. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50484/50484movie2.avi" target="_blank"> Click here to view movie</a>.
 Video 3. Video of the autophagosome formation event presented in Figure 4. The playback rate is 4 frames per second. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50484/50484movie3.avi" target="_blank"> Click here to view movie</a>. 
 
<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>DMEM</td>
 <td>Invitrogen</td>
 <td>41965</td>
 <td></td>
 </tr>
 <tr>
 <td>OptiMEM I</td>
 <td>Invitrogen</td>
 <td>31985-062</td>
 <td></td>
 </tr>
 <tr>
 <td>MitoTracker Red FM</td>
 <td>Invitrogen</td>
 <td>M22425</td>
 <td></td>
 </tr>
 <tr>
 <td>LysoTracker Red DND-99</td>
 <td>Invitrogen</td>
 <td>L-7528</td>
 <td></td>
 </tr>
 <tr>
 <td>X-tremeGENE 9 DNA Transfection Reagent</td>
 <td>Roche Applied Science</td>
 <td>6365787001</td>
 <td></td>
 </tr>
 <tr>
 <td>22 mm coverslips</td>
 <td>VWR</td>
 <td>631-0159</td>
 <td></td>
 </tr>
 <tr>
 <td>35 mm plates</td>
 <td>Fisher NUNC</td>
 <td>153066</td>
 <td></td>
 </tr>
 <tr>
 <td>Silicon grease</td>
 <td>RS Components Ltd. </td>
 <td>RS 494-124</td>
 <td></td>
 </tr>
 <tr>
 <td>O-rings</td>
 <td></td>
 <td></td>
 <td>Custom made</td>
 </tr>
 <tr>
 <td>Attofluor Cell Chamber</td>
 <td>Invitrogen</td>
 <td>A-7816</td>
 <td>Suggested alternative to custom-made O-rings</td>
 </tr>
 <tr>
 <td>Microscope</td>
 <td>Olympus</td>
 <td>IX81</td>
 <td>Inverted microscope</td>
 </tr>
 <tr>
 <td>Objective</td>
 <td>Olympus</td>
 <td>UPLSAPO 100XO</td>
 <td>N.A. 1.4, W.D. 0.13, FN 26.5</td>
 </tr>
 <tr>
 <td>Camera</td>
 <td>Hamamatsu</td>
 <td>ORCA-R2 C10600 10B</td>
 <td>Progressive scan interline CCD</td>
 </tr>
 <tr>
 <td>Illuminator</td>
 <td>TILL Photonics</td>
 <td>Polychrome V</td>
 <td>Ultrafast monochromator</td>
 </tr>
 <tr>
 <td>Incubation chamber</td>
 <td>Solent Scientific</td>
 <td>Cell^R IX81</td>
 <td></td>
 </tr>
 <tr>
 <td>Software</td>
 <td>Olympus</td>
 <td>SIS xcellence</td>
 <td></td>
 </tr>
 </tbody>
 </table>
Table 1. List of specific reagents and equipment required for the protocol, along with the corresponding provider and catalogue number. 
 Buffers 
 <table border="1">
 <tbody>
 <tr>
 <td fo:width=".5in">Buffer</td>
 <td fo:width="1.5in">Composition</td>
 <td fo:width=".5in">Step Used</td>
 </tr>
 <tr>
 <td>Starvation medium</td>
 <td>20 mM HEPES pH 7.4 </td>
 <td>5.2</td>
 </tr>
 <tr>
 <td></td>
 <td>140 mM NaCl </td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td>1 mM CaCl2 </td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td>1 mM MgCl2 </td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td>5 mM Glucose </td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td>1% BSA</td>
 <td></td>
 </tr>
 </tbody>
 </table>
 Table 2. List of buffers used in this protocol. The buffers used, their composition and the first step at which they are used in the protocol are listed.

Watch this Scientific Journal Video about Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond at JoVE.com

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

live-cell-imaging-of-early-autophagy-events-omegasomes-and-beyond

Nanyang Technological University

Research

JoVE Journal

Biology

18.4K Views.  The Babraham Institute. Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Video: Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Activating Autophagy by Aerobic Exercise in Mice

Autophagy is a lysosomal degradation pathway essential for cell homeostasis, function and differentiation. Under stress conditions, autophagy is induced and targets various cargos, such as bulk cytosol, damaged organelles and misfolded proteins, for degradation in lysosomes. Resulting nutrient molecules are recycled back to the cytosol for new protein synthesis and ATP production. Upregulation of autophagy has beneficial effects against the pathogenesis of many diseases, and pharmacological and physiological strategies to activate autophagy have been reported. Aerobic exercise is recently identified as an efficient autophagy inducer in multiple organs in mice, including muscle, liver, heart and brain. Here we show procedures to induce autophagy in vivo by either forced treadmill exercise or voluntary wheel running. We also demonstrate microscopic and biochemical methods to quantitatively analyze autophagy levels in mouse tissues, using the marker proteins LC3 and p62 that are transported to and degraded in lysosomes along with autophagosomes.

Autophagy activation is beneficial in the prevention of a number of diseases. One of the physiological approaches to induce autophagy in vivo is physical exercise. Here we show how to activate autophagy by aerobic exercise and measure autophagy levels in mice.

Autophagy is a lysosomal degradation pathway essential for cell homeostasis, function and differentiation. Under stress conditions, autophagy is induced ...

Live-cell Imaging of Platelet Degranulation and Secretion Under Flow

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the formation of thrombi that occlude the vasculature and injure organs, with life-threatening consequences. This motivates scientific research on platelet function and the development of methods to track cell-biological processes as they occur under flow conditions.
A variety of flow models are available for the study of platelet adhesion and aggregation, two key phenomena in platelet biology. This work describes a method to study real-time platelet degranulation under flow during activation. The method makes use of a flow chamber coupled to a syringe-pump setup that is placed under a wide-field, inverted, LED-based fluorescence microscope. The setup described here allows for the simultaneous excitation of multiple fluorophores that are delivered by fluorescently labeled antibodies or fluorescent dyes. After live-cell imaging experiments, the cover glasses can be further processed and analyzed using static microscopy (i.e., confocal microscopy or scanning electron microscopy).

This work describes a fluorescence microscopy-based method for the study of platelet adhesion, spreading, and secretion under flow. This versatile platform enables the investigation of platelet function for mechanistic research on thrombosis and hemostasis.

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the ...

LDL Cholesterol Uptake Assay Using Live Cell Imaging Analysis with Cell Health Monitoring

The regulation of LDL cholesterol uptake through LDLR-mediated endocytosis is an important area of study in various major pathologies including metabolic disorder, cardiovascular disease, and kidney disease. Currently, there is no available method to assess LDL uptake while simultaneously monitoring for health of the cells. The current study presents a protocol, using a live cell imaging analysis system, to acquire serial measurements of LDL influx with concurrent monitoring for cell health. This novel technique is tested in three human cell lines (hepatic, renal tubular epithelial, and coronary artery endothelial cells) over a four-hour time course. Moreover, the sensitivity of this technique is validated with well-known LDL uptake inhibitors, Dynasore and recombinant PCSK9 protein, as well as by an LDL uptake promoter, Simvastatin. Taken together, this method provides a medium-to-high throughput platform for simultaneously screening pharmacological activity as well as monitoring of cell morphology, hence cytotoxicity of compounds regulating LDL influx. The analysis can be used with different imaging systems and analytical software.

This protocol provides an efficient approach to measuring LDL cholesterol uptake with real time influx rates using a live cell imaging system in various cell types. This technique provides a platform to screen the pharmacological activity of compounds affecting LDL influx while monitoring for cell morphology and hence potential cytotoxicity.

The regulation of LDL cholesterol uptake through LDLR-mediated endocytosis is an important area of study in various major pathologies including metabolic ...