Time-lapse microscopy experiments were performed at the Biomolecular Screening Facility (BSF), EPFL. We thank Marc Delachaux (Service Audiovisuel, EPFL) for the videography and editing of the movie.

Note: In this study, the E14 ES cell line was used. However, this protocol is directly applicable to any other mouse ES cell line expressing a protein of interest fused to a SNAP-tag, either by tagging the endogenous protein or by using overexpression. For the examples shown in the results section, doxycycline-inducible SNAP-tag fusion cell lines were used (SNAP-tag fused to the following proteins: Nanog, Oct4, Srsf11, or to the fluorescent proteins mOrange2 and sfGFP, and put under the control of a doxycycline-inducible promoter. See11 for further information on the plasmids used for the generation of the doxycycline-inducible SNAP-tag fusion cell lines). The doxycycline-inducible system can be particularly useful, as it allows for tightly controlling the timing and intensity of the expression of the protein of interest. C-terminal positioning of the SNAP-tag is recommended, as changing the N-terminal amino acid sequence is more likely to alter the half-life of the target protein (N-end rule12).
1. E-cadherin coating and cell seeding
<ol>
	<li>Generate an ES cell line expressing a protein of interest fused to the SNAP-tag4.</li>
	<li>Coat 100 mm cell culture dishes with 4 mL of 0.1% gelatin (diluted in phosphate buffered saline (PBS) without Ca2+/Mg2+) for 1 h. Remove gelatin and let dry for 1 h. Use immediately or store the coated dishes for up to 2 months.</li>
	<li>Grow the cell line of interest in ES cell culture medium (Glasgow Minimum Essential Medium, supplemented with 10% ES cell-qualified fetal bovine serum, 2 mM sodium pyruvate, 1% non-essential amino acids, 1% penicillin/streptomycin, 2 mM L-glutamine, 100 &#181;M 2-mercaptoethanol, leukemia inhibitory factor (LIF), 3 &#181;M CHIR99021 and 0.8 &#181;M PD184352) on gelatin-coated dishes at 37 &#176;C and 5% CO2 for 2 days until reaching a confluence of 10 - 20 Mio cells per dish. 
	Note: Here, LIF was produced by transient transfection of HEK 293T cells, followed by supernatant collection and filtering. Each batch of LIF was tested for its potential to maintain pluripotency, but concentrations were not determined. However, recombinant LIF is also commercially available and the concentration commonly used for mouse ES cell culture is 1000 units/mL13.</li>
	<li>Coat a 96-well plate suitable for imaging with 30 &#181;L of recombinant mouse E-cadherin Fc chimera protein (E-cad-Fc) per well (5 ng/&#181;L, diluted in PBS with Ca2+/Mg2+. Use stock concentrations of 100 ng/&#181;L, stored at -80 &#176;C). Avoid extensive pipetting, as E-cad-Fc is very fragile. Incubate at 37 &#176;C for 1.5 h. 
	Note: Depending on the microscope, other formats might be used.</li>
	<li>Aspirate the E-cad-Fc, wash once with 100 &#181;L of PBS with Ca2+/Mg2+, and add 100 &#181;L of pre-warmed ES cell culture medium.</li>
	<li>Wash the cells of interest with 5 mL of PBS without Ca2+/Mg2+. Aspirate and add 2 mL of Trypsin-EDTA 0.25%. Incubate for 4 min at 37 &#176;C. Add 4 mL of ES cell culture medium, resuspend and spin down at 1000 x g for 4 min. Aspirate the supernatant and resuspend the pellet in 2 mL of fresh ES cell culture medium. 
	Note: PBS without Ca2+/Mg2+ should be used for washing the cells before trypsinization in order to remove Ca2+ ions, which are required for cell adhesion. In contrast, for the E-cad-Fc dilution and coating (step 1.4), use PBS with Ca2+/Mg2+, since the adhesion of ES cells to E-cad-Fc coated dishes is Ca2+-dependent14.</li>
	<li>Dilute the resuspended cells 1:10 in 1 mL of ES cell culture medium and count, using a counting chamber (load 10 &#181;L for a chamber of 0.1 mm depth).</li>
	<li>Seed 30,000 cells/cm2 in the E-cad-Fc coated imaging plate. Fill up to 200 &#181;L per well with ES cell culture medium. For doxycycline-inducible cell lines, induce with an appropriate dose of doxycycline (optimize dose and timing of the induction in advance. The cell lines used here were treated with 500 ng/mL doxycycline 24 h before imaging). Incubate at 37 &#176;C and 5% CO2 and proceed with the pulse labeling and imaging 24 h after cell seeding.</li>
</ol>
2. Pulse labeling of the SNAP-tag
Note: For protein decay experiments it is crucial to use an adequate SNAP dye concentration. The concentration should be high enough to yield a bright signal in the beginning of the time-lapse, as the fluorescence will decrease over time. However, using too high dye concentrations might cause residual dye being left in the medium or in the cells even after washing. The free dye might subsequently bind to newly produced SNAP-tag molecules over the course of the movie, which will distort the decay curve. The observed fluorescence signal will depend on the properties of the dye, the cell line used, as well as the expression level of the corresponding protein. Therefore, it is crucial to optimize the dye concentration by testing different dilutions, starting from the dilution suggested for live cell imaging by the manufacturer. For this study, a far-red fluorescent substrate was used. An optimal concentration of 12 nM was determined for doxycycline-inducible overexpression cell lines.
<ol>
	<li>Dilute the SNAP dye to the appropriate concentration in pre-warmed ES cell culture medium. Use a pipette to gently remove the medium from the cells previously seeded in the 96-well plate and add 50 &#181;L of diluted SNAP dye per well. Incubate for 30 min at 37 &#176;C.</li>
	<li>Aspirate the diluted dye with a pipette and wash 3x with 200 &#181;L of pre-warmed PBS (without Ca2+/Mg2+). Add 200 &#181;L of ES cell culture medium and incubate for 15 min at 37 &#176;C.</li>
	<li>Repeat the previous washing steps twice more. 
	Note: Extensive washing is important in order to remove any unbound dye. The repeated washing steps with PBS remove the residual extracellular dye in the medium, whereas the 15 min incubations ensure robust labeling of the SNAP-tag fusion proteins prior to starting the imaging experiment. However, perform the washing steps by gentle pipetting and do not use an automatic aspirator, as cells seeded on E-cad-Fc tend to detach easily.</li>
	<li>Add 200 &#181;L of imaging medium. To reduce the fluorescence background, use phenol red free medium (supplemented with 10% ES cell-qualified fetal bovine serum, 2 mM sodium pyruvate, 1% non-essential amino acids, 1% penicillin/streptomycin, 2 mM L-glutamine, 100 &#181;M 2-mercaptoethanol, LIF, 3 &#181;M CHIR99021 and 0.8 &#181;M PD184352).</li>
</ol>
3. Time-lapse microscopy
<ol>
	<li>Use a microscope suitable for live imaging, allowing controlled temperature and CO2. Set the temperature to 37 &#176;C and the CO2 to 5% prior to use and allow to equilibrate for 1 - 2 h.</li>
	<li>Place the plate into the microscope and find spots suitable for imaging. Select spots where the cells are evenly distributed, but not too dense, as this will facilitate the analysis. Adjust the number of selected spots to the number of cells required for the analysis. In this study, 3 -&#160;5 spots per cell line were recorded.</li>
	<li>Select the objective. A 20X objective is appropriate for the size of ES cells.</li>
	<li>Select the illumination settings for the fluorescent channel(s) to be recorded. Adjust the laser power and exposure according to the intensity of the fluorescence signal from the cells of interest. Make sure to obtain a strong initial signal, since it will decrease over the course of the movie, but avoid laser powers higher than 30% in order to minimize phototoxicity and photobleaching. For this study, an excitation filter of 632/22 nm (wavelength/bandpass), emission filter of 679/34 nm (wavelength/bandpass), laser power of 10% and exposure time of 100 ms were used.</li>
	<li>Select the imaging intervals and duration of the time-lapse experiment. For proteins with expected half-lives of 2 - 20 h, choose acquisition times of 12 - 24 h with time intervals of 15 min. For shorter-lived proteins, reduce the intervals and acquisition times, for longer-lived proteins increase accordingly.</li>
	<li>Start imaging.</li>
</ol>
4. Image processing and analysis
<ol>
	<li>Collect the acquired images as a stack in tiff format. For further processing and analysis, use the FIJI software15. If the time-lapse data is not collected as a stack by the microscope software, open all frames of the time-lapse experiment in the software (either by clicking File | Open&#8230; or drag and dropping the corresponding files to the toolbar) and click Image | Stacks | Images to Stack.</li>
	<li>Use the subtract background function to remove the background from all pictures in the stack. To do so, click Process | Subtract Background. Select the rolling ball radius in the pop-up window. Use a radius that is at least the size of the cells imaged. To apply the background subtraction to all cells in the stack, select Yes in the Process stack? pop-up window.</li>
	<li>Select a cell of interest and draw a region of interest (ROI) around it using the toolbar. For nuclear signals, use the Oval selection by first clicking on the corresponding tab in the toolbar and then clicking on the nucleus of interest and adjusting the oval around it. For cytoplasmic signals or very dense regions use the freehand selection to be able to closely follow the cell outlines. Avoid including too large regions of background, even though it is not necessary to follow the cell outlines precisely due to the subsequent subtraction of all background pixels (step 4.8 - 4.9).</li>
	<li>Add the region of interest to the ROI manager by clicking Analyze | Tools | ROI manager | Add, or by pressing T on the keyboard.</li>
	<li>Proceed to the next frame by moving the tab in the stack window or by pressing shift + &#60; on the keyboard, then repeat the previous actions. Follow the cell of interest throughout the course of the movie and add the ROI of each frame to the ROI manager. Save the final ROI set for each tracked cell by clicking More | Save&#8230; in the ROI manager. 
	Note: After cell divisions, track both daughter cells separately and sum their integrated intensities after background subtraction (see steps 4.6 - 4.9) to account for protein dilution during cell division. Save the ROI sets for both daughter cells.</li>
	<li>Once the cell of interest is tracked throughout the movie, click Measure, to obtain values for the mean intensity and the area of the ROIs. Copy the obtained values into an electronic spreadsheet program and calculate the raw integrated intensity of the cell for each time-point as follows: 
	<img alt="Equation 1" src="/files/ftp_upload/56604/56604eq1.jpg" /> 
	where Meanc is the mean intensity and AreaC the area of the corresponding ROI.</li>
	<li>To estimate the local background, add a ROI close to the cell of interest for each time frame. Avoid including any cellular fluorescence signal. Use a circular ROI that is roughly the size of a cell and move or shrink it accordingly if needed, e.g. if neighboring cells interfere. Proceed as described previously with the cellular ROI set in order to obtain a background ROI set and copy the measured intensity values to the spreadsheet.</li>
	<li>To obtain a background-corrected value for the integrated intensity of the cell, first calculate the integrated intensity of the background for each time-point: 
	<img alt="Equation 2" src="/files/ftp_upload/56604/56604eq2.jpg" /> 
	where MeanBG is the mean intensity of the background signal and AreaC is the area of the ROI encircling the cell. Do not use the area of the background ROI, unless both areas have the same size.</li>
	<li>Calculate the final background-subtracted integrated intensity of the cell for each time point: 
	<img alt="Equation 3" src="/files/ftp_upload/56604/56604eq3.jpg" /></li>
	<li>To normalize the single cell decay curves, divide the intensity value of each time-point by the intensity value of the first time-point. 
	Note: The curve fitting (see step 4.11) can either be performed on each single cell or on the population average. A normalization is required if a population based average is calculated in order to avoid biases from different fluorescence intensities between cells. The normalization thus ensures that each cell contributes with the same weight to the final decay curve. In addition, the normalization can be useful to visualize the single cell decays independently of their absolute fluorescence intensities (see Figures 3A-3C). However, if only the single-cell half-lives are determined and the data is not averaged, the normalization step may be omitted and the curve fitting can be directly performed on the raw data.</li>
	<li>For the estimation of the protein half-life, use a curve fitting tool. In this study, the MATLAB curve fitting toolbox 3.4.1 was used, which is located by default in the Apps section of the MATLAB user interface. Import the fluorescence intensity and time values from the electronic spreadsheet into MATLAB by clicking on the Import Data tab. Open the curve fitting toolbox and select the time points and fluorescence decay data in the X data and Y data tab. Choose Custom Equation in the curve fitting tab and enter the equation for an exponential decay: 
	<img alt="Equation 4" src="/files/ftp_upload/56604/56604eq4.jpg" /> 
	where f(t) is the fluorescence intensity at a given time-point, a the initial intensity and b the decay rate. In the Fit Options&#8230; tab, select 0 for the lower limit of both a and b. The estimated values for a and b will then appear in the results window. Calculate the half-life as follows: 
	<img alt="Equation 5" src="/files/ftp_upload/56604/56604eq5.jpg" /></li>
</ol>

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The authors have nothing to disclose.

The most crucial step when using a SNAP-tag to monitor protein decay is to ensure that no residual unbound dye is left in the medium or in the cells after washing, as otherwise it might bind to newly produced SNAP-tag molecules later in the course of the experiment and thereby compromise the decay curve. This is on one hand achieved by carefully performing all the described washing steps. On the other hand, the dye concentration should be kept as low as possible, while still being in a range that allows for obtaining a g...

It is well known that cellular proteins undergo extensive turnover, with synthesis and degradation rates being specific for each protein and subject to physiological regulation. Traditionally, protein degradation rates have been measured using bulk methods, such as radioactive pulse chase analysis, or involving protein synthesis inhibitors such as cycloheximide1. More recently, stable isotope labeling with amino acids in cell culture (SILAC) in combination with mass spectrometry has been established to quantify protein turnover on a global scale2. However, these methods are limited by population averaging, and information about cell-to-cell variability is therefore lost. Furthermore, transient changes in protein degradation that are unsynchronized across the cell population cannot be identified.
Alternatively, protein half-lives can also be determined by fluorescence-based approaches, which often have the advantage of providing single-cell resolution. For example, a photoactivatable green fluorescent protein (paGFP) has been used to determine Oct4 half-life in the early mammalian embryo3. Another method to monitor protein decay in living cells is the use of a SNAP-tag in combination with fluorescence time-lapse imaging. The SNAP-tag is a mutant version of the DNA repair enzyme O6-alkylguanine DNA-alkyltransferase (AGT) that specifically reacts with benzylguanine (BG) derivatives, which can be coupled to molecular probes4,5,6. Therefore, any SNAP-tag fusion protein can be irreversibly labeled with a fluorescent, cell permeable dye. Pulse labeling of a protein of interest fused to the SNAP-tag, followed by washout of the residual dye, allows for monitoring the degradation of the labeled protein population and thus for determining protein half-life. SNAP-tags have been successfully used for pulse-chase labeling of proteins and for determining protein half-lives in adherent cell culture and in vivo5,7,8,9. A large variety of SNAP-tag substrates covering commonly used fluorescent spectra are commercially available, enabling the selection of the optimal dye for each specific application. Thus, SNAP-tags can also be used for multicolor imaging in combination with other fluorescent fusion proteins or dyes. Cell-impermeable dyes are suitable for labeling of membrane-tethered proteins, whereas cell-permeable dyes are applicable for monitoring both intracellular and membrane-bound proteins. Furthermore, some of these probes exhibit almost no basal fluorescence and only start emitting a strong fluorescent signal upon binding to a SNAP-tag10.
This protocol describes how to measure the degradation rates of different proteins of interest in single cells using a SNAP-tag. Here we apply this method to mouse embryonic stem (ES) cells cultured on E-cadherin, but it should be possible to use it with any adherent cultured cell type. We show that pulse labeling of SNAP-tag fusion proteins followed by fluorescence time-lapse imaging allows for determining the single cell half-lives of various proteins of interest and provides an estimate for the cell-to-cell variability of half-lives in a population of cultured cells.

<table border="0" cellpadding="0" cellspacing="0" width="803">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Equipment</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:329px;">ES cell line expressing a SNAP-tag fusion protein of interest</td>
			<td style="width:189px;">-</td>
			<td style="width:119px;">-</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Falcon 100 mm TC-Treated Cell Culture Dish</td>
			<td style="width:189px;">Corning</td>
			<td style="width:119px;">353003</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:329px;">96 Well, Black/Clear, Tissue Culture Treated Plate</td>
			<td style="width:189px;">Corning</td>
			<td style="width:119px;">353219</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Neubauer-improved counting chamber, 0.1 mm</td>
			<td style="width:189px;">Marienfeld-superior</td>
			<td style="width:119px;">640030</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:329px;">CO2 Incubator</td>
			<td style="width:189px;">Panasonic</td>
			<td style="width:119px;">MCO-170AICUV-PE</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Centrifuge 5804 R</td>
			<td style="width:189px;">Eppendorf</td>
			<td>5804000528</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:329px;">InCell Analyzer 2200 Cell Imaging System</td>
			<td style="width:189px;">GE Healthcare Life Sciences</td>
			<td style="width:119px;">29027886</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Reagents</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Glasgow Minimum Essential Medium</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td>G5154</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:329px;">Fetal Bovine Serum, embyonic stem cell-qualified</td>
			<td style="width:189px;">ThermoFisher</td>
			<td style="width:119px;">16141-079</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Sodium pyruvate solution</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">113-24-6</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:329px;">Minimum Essential Medium&#160; Non-Essential Amino Acids</td>
			<td style="width:189px;">ThermoFisher</td>
			<td style="width:119px;">11140-035</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Penicillin-Streptomycin</td>
			<td style="width:189px;">BioConcept</td>
			<td style="width:119px;">4-01F00H</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">L-Glutamine 200mM</td>
			<td style="width:189px;">ThermoFisher</td>
			<td style="width:119px;">25030-024</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">2-Mercaptoethanol</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">63689-25ML-F</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Leukemia Inhibitory factor</td>
			<td style="width:189px;">-</td>
			<td style="width:119px;">-</td>
			<td>Produced in the lab by transient transfection of HEK-293T cells, followed by collection and filtering of the supernatant.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">CHIR99021 (GSK-3 Inhibitor XVI)</td>
			<td style="width:189px;">Merck Millipore</td>
			<td style="width:119px;">361559</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">PD 0325901</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">391210-10-9</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Gelatin from bovine skin</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">9000-70-8</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Dulbecco&#39;s PBS 10x concentrated</td>
			<td style="width:189px;">BioConcept</td>
			<td style="width:119px;">3-05K00-I</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Dulbecco&#39;s PBS Without Ca++/Mg++</td>
			<td style="width:189px;">BioConcept</td>
			<td style="width:119px;">3-05F29-I</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Trypsin-EDTA-Solution 0.25%</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">T4049</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:329px;">Recombinant Mouse E-Cadherin Fc Chimera protein</td>
			<td style="width:189px;">R&#38;D systems</td>
			<td style="width:119px;">748-EC-050</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Doxycycline hyclate</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">D9891</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">SNAP-Cell 647-SiR</td>
			<td style="width:189px;">New England BioLabs</td>
			<td style="width:119px;">S9102S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">FluoroBrite DMEM</td>
			<td style="width:189px;">ThermoFisher</td>
			<td style="width:119px;">A18967-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Software</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">FIJI</td>
			<td style="width:189px;">-</td>
			<td style="width:119px;">-</td>
			<td>Open-source image analysis software</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">MATLAB R2014a</td>
			<td style="width:189px;">Mathworks</td>
			<td style="width:119px;">-</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:329px;">Microsoft Excel</td>
			<td style="width:189px;">Microsoft</td>
			<td style="width:119px;">-</td>
			<td></td>
		</tr>
	</tbody>
</table>

single-cell quantification of protein degradation rates by time-lapse fluorescence microscopy in adherent cell culture

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used approaches to determine protein half-life are either limited to population averages from lysed cells or require the use of protein synthesis inhibitors. This protocol describes a method to measure protein half-lives in single living adherent cells, using SNAP-tag fusion proteins in combination with fluorescence time-lapse microscopy. Any protein of interest fused to a SNAP-tag can be covalently bound by a fluorescent, cell permeable dye that is coupled to a benzylguanine derivative, and the decay of the labeled protein population can be monitored after washout of the residual dye. Subsequent cell tracking and quantification of the integrated fluorescence intensity over time results in an exponential decay curve for each tracked cell, allowing for determining protein degradation rates in single cells by curve fitting. This method provides an estimate for the heterogeneity of half-lives in a population of cultured cells, which cannot easily be assessed by other methods. The approach presented here is applicable to any type of cultured adherent cells expressing a protein of interest fused to a SNAP-tag. Here we use mouse embryonic stem (ES) cells grown on E-cadherin-coated cell culture plates to illustrate how single cell degradation rates of proteins with a broad range of half-lives can be determined.

The described protocol provides an estimate of the cell-to-cell variability in half-life for any given protein fused to a SNAP-tag. The use of recombinant E-cadherin-Fc for coating of the imaging plate allows for single cell resolution in ES cells, which otherwise grow in colonies. Single cells can be tracked separately throughout the course of the movie ( Figure 1A).
In order to determine the protein half-life for each single cell, it is crucial to measure the in...

Watch this Scientific Journal Video about Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture at JoVE.com

Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

This protocol describes a method to determine protein half-lives in single living adherent cells, using pulse labeling and fluorescence time-lapse imaging of SNAP-tag fusion proteins.

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used ...

single-cell-quantification-protein-degradation-rates-time-lapse

Research

JoVE Journal

Developmental Biology

8.5K Views.  École Polytechnique Fédérale de Lausanne (EPFL). This protocol describes a method to determine protein half-lives in single living adherent cells, using pulse labeling and fluorescence time-lapse imaging of SNAP-tag fusion proteins.

Video: Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Clonal Analysis of Embryonic Hematopoietic Stem Cell Precursors Using Single Cell Index Sorting Combined with Endothelial Cell Niche Co-culture

The ability to study hematopoietic stem cell (HSC) genesis during embryonic development has been limited by the rarity of HSC precursors in the early embryo and the lack of assays that functionally identify the long-term multilineage engraftment potential of individual putative HSC precursors. Here, we describe methodology that enables the isolation and characterization of functionally validated HSC precursors at the single cell level. First, we utilize index sorting to catalog the precise phenotypic parameter of each individually sorted cell, using a combination of phenotypic markers to enrich for HSC precursors with additional markers for experimental analysis. Second, each index-sorted cell is co-cultured with vascular niche stroma from the aorta-gonad-mesonephros (AGM) region, which supports the maturation of non-engrafting HSC precursors to functional HSC with multilineage, long-term engraftment potential in transplantation assays. This methodology enables correlation of phenotypic properties of clonal hemogenic precursors with their functional engraftment potential or other properties such as transcriptional profile, providing a means for the detailed analysis of HSC precursor development at the single cell level.

Here we present methodology for the clonal analysis of hematopoietic stem cell precursors during murine embryonic development. We combine index sorting of single cells from the embryonic aorta-gonad-mesonephros region with endothelial cell co-culture and transplantation to characterize the phenotypic properties and engraftment potential of single hematopoietic precursors.

The ability to study hematopoietic stem cell (HSC) genesis during embryonic development has been limited by the rarity of HSC precursors in the early ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Single-cell Transcriptomic Analyses of Mouse Pancreatic Endocrine Cells

Pancreatic endocrine cells, which are clustered in islets, regulate blood glucose stability and energy metabolism. The distinct cell types in islets, including insulin-secreting &#946; cells, are differentiated from common endocrine progenitors during the embryonic stage. Immature endocrine cells expand via cell proliferation and mature during a long postnatal developmental period. However, the mechanisms underlying these processes are not clearly defined. Single-cell RNA-sequencing is a promising approach for the characterization of distinct cell populations and tracing cell lineage differentiation pathways. Here, we describe a method for the single-cell RNA-sequencing of isolated pancreatic &#946; cells from embryonic, neonatal and postnatal pancreases.

We describe a method for the isolation of endocrine cells from embryonic, neonatal and postnatal pancreases followed by single-cell RNA sequencing. This method allows analyses of pancreatic endocrine lineage development, cell heterogeneity and transcriptomic dynamics.

Pancreatic endocrine cells, which are clustered in islets, regulate blood glucose stability and energy metabolism. The distinct cell types in islets, ...

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

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Rozdziały w tym wideo

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

Clonal Analysis of Embryonic Hematopoietic Stem Cell Precursors Using Single Cell Index Sorting Combined with Endothelial Cell Niche Co-culture

Single-cell Photoconversion in Living Intact Zebrafish

Single-cell Transcriptomic Analyses of Mouse Pancreatic Endocrine Cells

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos