Name: single-cell quantification of protein degradation rates by time-lapse fluorescence microscopy in adherent cell culture
Uploaded: 2018-02-04T15:00:00.000+00:00
Duration: 12 min 6 s
Description: 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

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><tbody><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>ES cell line expressing a SNAP-tag fusion protein of interest</td><td>-</td><td>-</td><td></td></tr><tr><td>Falcon 100 mm TC-Treated Cell Culture Dish</td><td>Corning</td><td>353003</td><td></td></tr><tr><td>96 Well, Black/Clear, Tissue Culture Treated Plate</td><td>Corning</td><td>353219</td><td></td></tr><tr><td>Neubauer-improved counting chamber, 0.1 mm</td><td>Marienfeld-superior</td><td>640030</td><td></td></tr><tr><td>CO2 Incubator</td><td>Panasonic</td><td>MCO-170AICUV-PE</td><td></td></tr><tr><td>Centrifuge 5804 R</td><td>Eppendorf</td><td>5804000528</td><td></td></tr><tr><td>InCell Analyzer 2200 Cell Imaging System</td><td>GE Healthcare Life Sciences</td><td>29027886</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Glasgow Minimum Essential Medium</td><td>Sigma-Aldrich</td><td>G5154</td><td></td></tr><tr><td>Fetal Bovine Serum, embyonic stem cell-qualified</td><td>ThermoFisher</td><td>16141-079</td><td></td></tr><tr><td>Sodium pyruvate solution</td><td>Sigma-Aldrich</td><td>113-24-6</td><td></td></tr><tr><td>Minimum Essential Medium&amp;nbsp; Non-Essential Amino Acids</td><td>ThermoFisher</td><td>11140-035</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>BioConcept</td><td>4-01F00H</td><td></td></tr><tr><td>L-Glutamine 200mM</td><td>ThermoFisher</td><td>25030-024</td><td></td></tr><tr><td>2-Mercaptoethanol</td><td>Sigma-Aldrich</td><td>63689-25ML-F</td><td></td></tr><tr><td>Leukemia Inhibitory factor</td><td>-</td><td>-</td><td>Produced in the lab by transient transfection of HEK-293T cells, followed by collection and filtering of the supernatant.&amp;nbsp;</td></tr><tr><td>CHIR99021 (GSK-3 Inhibitor XVI)</td><td>Merck Millipore</td><td>361559</td><td></td></tr><tr><td>PD 0325901</td><td>Sigma-Aldrich</td><td>391210-10-9</td><td></td></tr><tr><td>Gelatin from bovine skin</td><td>Sigma-Aldrich</td><td>9000-70-8</td><td></td></tr><tr><td>Dulbecco&#039;s PBS 10x concentrated</td><td>BioConcept</td><td>3-05K00-I</td><td></td></tr><tr><td>Dulbecco&#039;s PBS Without Ca++/Mg++</td><td>BioConcept</td><td>3-05F29-I</td><td></td></tr><tr><td>Trypsin-EDTA-Solution 0.25%</td><td>Sigma-Aldrich</td><td>T4049</td><td></td></tr><tr><td>Recombinant Mouse E-Cadherin Fc Chimera protein</td><td>R&amp;amp;D systems</td><td>748-EC-050</td><td></td></tr><tr><td>Doxycycline hyclate</td><td>Sigma-Aldrich</td><td>D9891</td><td></td></tr><tr><td>SNAP-Cell 647-SiR</td><td>New England BioLabs</td><td>S9102S</td><td></td></tr><tr><td>FluoroBrite DMEM</td><td>ThermoFisher</td><td>A18967-01</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>FIJI</td><td>-</td><td>-</td><td>Open-source image analysis software</td></tr><tr><td>MATLAB R2014a</td><td>Mathworks</td><td>-</td><td></td></tr><tr><td>Microsoft Excel</td><td>Microsoft</td><td>-</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

8.6K 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

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Measurement of Protein Turnover Rates in Senescent and Non-Dividing Cultured Cells with Metabolic Labeling and Mass Spectrometry

Mounting evidence has shown that the accumulation of senescent cells in the central nervous system contributes to neurodegenerative disorders such as Alzheimer&#39;s and Parkinson&#39;s diseases. Cellular senescence is a state of permanent cell cycle arrest that typically occurs in response to exposure to sub-lethal stresses. However, like other non-dividing cells, senescent cells remain metabolically active and carry out many functions that require unique transcriptional and translational demands and widespread changes in the intracellular and secreted proteomes. Understanding how protein synthesis and decay rates change during senescence can illuminate the underlying mechanisms of cellular senescence and find potential therapeutic avenues for diseases exacerbated by senescent cells. This paper describes a method for proteome-scale evaluation of protein half-lives in non-dividing cells using pulsed stable isotope labeling by amino acids in cell culture (pSILAC) in combination with mass spectrometry. pSILAC involves metabolic labeling of cells with stable heavy isotope-containing versions of amino acids. Coupled with modern mass spectrometry approaches, pSILAC enables the measurement of protein turnover of hundreds or thousands of proteins in complex mixtures. After metabolic labeling, the turnover dynamics of proteins can be determined based on the relative enrichment of heavy isotopes in peptides detected by mass spectrometry. In this protocol, a workflow is described for the generation of senescent fibroblast cultures and similarly arrested quiescent fibroblasts, as well as a simplified, single-time point pSILAC labeling time-course that maximizes coverage of anticipated protein turnover rates. Further, a pipeline is presented for the analysis of pSILAC mass spectrometry data and user-friendly calculation of protein degradation rates using spreadsheets. The application of this protocol can be extended beyond senescent cells to any non-dividing cultured cells such as neurons.

This protocol describes the workflow for metabolic labeling of senescent and non-dividing cells with pulsed SILAC, untargeted mass spectrometry analysis, and a streamlined calculation of protein half-lives.

Mounting evidence has shown that the accumulation of senescent cells in the central nervous system contributes to neurodegenerative disorders such as ...

Biochemistry

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Biology

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

Immunology and Infection

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts 

Cellular invasion into local tissues is a process important in development and homeostasis. Malregulated invasion and subsequent cell movement is characteristic of multiple pathological processes, including inflammation, cardiovascular disease and tumor cell metastasis1. Focalized proteolytic degradation of extracellular matrix (ECM) components in the epithelial or endothelial basement membrane is a critical step in initiating cellular invasion. In tumor cells, extensive in vitro analysis has determined that ECM degradation is accomplished by ventral actin-rich membrane protrusive structures termed invadopodia2,3. Invadopodia form in close apposition to the ECM, where they moderate ECM breakdown through the action of matrix metalloproteinases (MMPs). The ability of tumor cells to form invadopodia directly correlates with the ability to invade into local stroma and associated vascular components3. 
Visualization of invadopodia-mediated ECM degradation of cells by fluorescent microscopy using dye-labeled matrix proteins coated onto glass coverslips has emerged as the most prevalent technique for evaluating the degree of matrix proteolysis and cellular invasive potential4,5. Here we describe a version of the standard method for generating fluorescently-labeled glass coverslips utilizing a commercially available Oregon Green-488 gelatin conjugate. This method is easily scaled to rapidly produce large numbers of coated coverslips. We show some of the common microscopic artifacts that are often encountered during this procedure and how these can be avoided. Finally, we describe standardized methods using readily available computer software to allow quantification of labeled gelatin matrix degradation mediated by individual cells and by entire cellular populations. The described procedures provide the ability to accurately and reproducibly monitor invadopodia activity, and can also serve as a platform for evaluating the efficacy of modulating protein expression or testing of anti-invasive compounds on extracellular matrix degradation in single and multicellular settings.

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

We describe the prototypical method for producing microscope coverslips coated with fluorescent gelatin for visualizing invadopodia-mediated matrix degradation. Computational techniques using available software are presented for quantifying the resultant levels of matrix proteolysis by single cells within a mixed population and for multicellular groups encompassing entire microscopic fields.

Cellular invasion into local tissues is a process important in development and homeostasis.  Malregulated invasion and subsequent cell movement is ...

Temporal Quantification of MAPK Induced Expression in Single Yeast Cells

The quantification of gene expression at the single cell level uncovers novel regulatory mechanisms obscured in measurements performed at the population level. Two methods based on microscopy and flow cytometry are presented to demonstrate how such data can be acquired. The expression of a fluorescent reporter induced upon activation of the high osmolarity glycerol MAPK pathway in yeast is used as an example. The specific advantages of each method are highlighted. Flow cytometry measures a large number of cells (10,000) and provides a direct measure of the dynamics of protein expression independent of the slow maturation kinetics of the fluorescent protein. Imaging of living cells by microscopy is by contrast limited to the measurement of the matured form of the reporter in fewer cells. However, the data sets generated by this technique can be extremely rich thanks to the combinations of multiple reporters and to the spatial and temporal information obtained from individual cells. The combination of these two measurement methods can deliver new insights on the regulation of protein expression by signaling pathways.

Two complementary methods based on flow cytometry and microscopy are presented which enable the quantification, at the single cell level, of the dynamics of gene expression induced by the activation of a MAPK pathway in yeast.

The quantification of gene expression at the  single cell level uncovers novel regulatory mechanisms obscured in measurements  performed at the population ...

Reporter-based Growth Assay for Systematic Analysis of Protein Degradation

Protein degradation by the ubiquitin-proteasome system (UPS) is a major regulatory mechanism for protein homeostasis in all eukaryotes. The standard approach to determining intracellular protein degradation relies on biochemical assays for following the kinetics of protein decline. Such methods are often laborious and time consuming and therefore not amenable to experiments aimed at assessing multiple substrates and degradation conditions. As an alternative, cell growth-based assays have been developed, that are, in their conventional format, end-point assays that cannot quantitatively determine relative changes in protein levels.
Here we describe a method that faithfully determines changes in protein degradation rates by coupling them to yeast cell-growth kinetics. The method is based on an established selection system where uracil auxotrophy of URA3-deleted yeast cells is rescued by an exogenously expressed reporter protein, comprised of a fusion between the essential URA3 gene and a degradation determinant (degron). The reporter protein is designed so that its synthesis rate is constant whilst its degradation rate is determined by the degron. As cell growth in uracil-deficient medium is proportional to the relative levels of Ura3, growth kinetics are entirely dependent on the reporter protein degradation.
This method accurately measures changes in intracellular protein degradation kinetics. It was applied to: (a) Assessing the relative contribution of known ubiquitin-conjugating factors to proteolysis (b) E2 conjugating enzyme structure-function analyses (c) Identification and characterization of novel degrons. Application of the degron-URA3-based system transcends the protein degradation field, as it can also be adapted to monitoring changes of protein levels associated with functions of other cellular pathways.

Here we describe a robust biological assay for quantifying the relative rate of proteolysis by the ubiquitin-proteasome system. The assay readout is yeast growth rate in liquid culture, which is dependent on the cellular levels of a reporter protein comprising a degradation signal fused to an essential metabolic marker.

Protein degradation by the ubiquitin-proteasome system (UPS) is a major regulatory mechanism for protein homeostasis in all eukaryotes. The standard ...

Assays for the Degradation of Misfolded Proteins in Cells

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded proteins may serve as potential therapeutic targets. To distinguish degradation of misfolding-prone proteins from other mechanisms that regulate their levels, one important method is to measure protein half-life in cells. However, this can be challenging because misfolding-prone proteins may exist in different forms, including the native form and misfolded forms of distinct characteristics. Here we describe assays to examine the half-life of misfolded proteins in mammalian cells using a highly aggregation-prone protein, Ataxin-1 with an extended polyglutamine (polyQ) stretch, and a conformationally unstable luciferase mutant as models. Cycloheximide chase is combined with cell fractionation to examine the turnover rate of misfolding-prone proteins in various cellular fractions. We further depict a fluorescence-based assay using an enhanced green fluorescence protein (EGFP)-fusion of the luciferase mutant, which can be adapted for high throughput screening on a microplate-reader.

This report describes protocols for measuring degradation rates of misfolded proteins by either western blot or fluorescence-based assays. The methods can be applied to analysis of other misfolded proteins and for high throughput screening.

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded ...

Measuring Diurnal Rhythms in Autophagic and Proteasomal Flux

Cells employ several methods for recycling unwanted proteins and other material, including lysosomal and non-lysosomal pathways. The main lysosome-dependent pathway is called autophagy, while the primary non-lysosomal method for protein catabolism is the ubiquitin-proteasome system. Recent studies in model organisms suggest that the activity of both autophagy and the ubiquitin-proteasome system is not constant across the day but instead varies according to a daily (circadian) rhythm. The ability to measure biological rhythms in protein turnover is important for understanding how cellular quality control is achieved and for understanding the dynamics of specific proteins of interest. Here we present a standardized protocol for quantifying autophagic and proteasomal flux in vivo that captures the circadian component of protein turnover. Our protocol includes details for mouse handling, tissue processing, fractionation, and autophagic flux quantification using mouse liver as the starting material.

We describe our protocol for measuring biological rhythms in protein catabolism via autophagy and the proteasome in mouse liver.

Cells employ several methods for recycling unwanted proteins and other material, including lysosomal and non-lysosomal pathways. The main ...

Live-cell Imaging of Single-Cell Arrays (LISCA) - a Versatile Technique to Quantify Cellular Kinetics

Live-cell Imaging of Single-Cell Arrays (LISCA) is a versatile method to collect time courses of fluorescence signals from individual cells in high throughput. In general, the acquisition of single-cell time courses from cultured cells is hampered by cell motility and diversity of cell shapes. Adhesive micro-arrays standardize single-cell conditions and facilitate image analysis. LISCA combines single-cell microarrays with scanning time-lapse microscopy and automated image processing. Here, we describe the experimental steps of taking single-cell fluorescence time courses in a LISCA format. We transfect cells adherent to a micropatterned array using mRNA encoding for enhanced green fluorescent protein (eGFP) and monitor the eGFP expression kinetics of hundreds of cells in parallel via scanning time-lapse microscopy. The image data stacks are automatically processed by newly developed software that integrates fluorescence intensity over selected cell contours to generate single-cell fluorescence time courses. We demonstrate that eGFP expression time courses after mRNA transfection are well described by a simple kinetic translation model that reveals expression and degradation rates of mRNA. Further applications of LISCA for event time correlations of multiple markers in the context of signaling apoptosis are discussed.

Live-cell Imaging of Single-Cell Arrays LISCA - a Versatile Technique to Quantify Cellular Kinetics

We present a method for the acquisition of fluorescence reporter time courses from single cells using micropatterned arrays. The protocol describes the preparation of single-cell arrays, the setup and operation of live-cell scanning time-lapse microscopy and an open-source image analysis tool for automated preselection, visual control and tracking of cell-integrated fluorescence time courses per adhesion site.

Live-cell Imaging of Single-Cell Arrays (LISCA) is a versatile method to collect time courses of fluorescence signals from individual cells in high ...

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

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Chapters in this video

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Measurement of Protein Turnover Rates in Senescent and Non-Dividing Cultured Cells with Metabolic Labeling and Mass Spectrometry

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Temporal Quantification of MAPK Induced Expression in Single Yeast Cells

Reporter-based Growth Assay for Systematic Analysis of Protein Degradation

Assays for the Degradation of Misfolded Proteins in Cells

Measuring Diurnal Rhythms in Autophagic and Proteasomal Flux

Live-cell Imaging of Single-Cell Arrays (LISCA) - a Versatile Technique to Quantify Cellular Kinetics