We are grateful to S. Taylor for providing the pLPCX-Histone H2-GFP plasmid. We thank R. Mertelsmann for continuous support. This work was supported by the Deutsche Forschungsgemeinschaft.

1. Seeding of U2OS-based Cyclin B-SNAP Reporter Cells (Clone 11 Cells 6) on Microscope Chamber Slides
<ol>
	<li>Trypsinize subconfluent SNAP reporter cells that were allowed to grow asynchronously in log phase for at least 48 hr.</li>
	<li>Working with 8 well microscope chambers (constant distribution of cells).</li>
</ol>
For seeding of cells onto 8 well microscope chambers at a constant distribution across the entire surface of the microscope chamber, centrifuge 10,000 cells and resuspend in 350 &mu;l of phenol red-free normal growth medium (supplemented with 10% fetal bovine serum, penicillin/streptomycin and sodium pyruvate). Transfer cell suspension to the microscope chamber (Figure 2).
Working with 8 well microscope chambers (maximum cell density in the center).
For seeding of cells at a higher density in the center of the microscope chamber, load the chamber with 300 &mu;l of phenol red-free normal growth medium. Add 5,000 cells carefully to the center of the microscope chamber (Figure 2).
Working with 96 well special optics plates (constant distribution of cells). 
For seeding of cells onto 96 well plates at a constant distribution across the entire surface of the well, centrifuge 5,000 cells and resuspend in 300 &mu;l of phenol red-free normal growth medium. Transfer the cell suspension to a 96-well plate. Depending on the total number of cell-containing wells required, adjust the cell number and the total volume of the suspension medium (Figure 2).
Working with 96 well special optics plates (maximum cell density in the center).
For seeding of cells onto 96 well plates, carefully add 1,500 cells in 15 &mu;l of phenol red-free normal growth medium in a small drop to the center of each well to achieve restriction of cell growth to the center of the well (Figure 2).
<ol start="3">
	<li>Allow seeded cells to grow for at least 18 hr under standard cell culture conditions (37 &deg;C, 100% air humidity, 5% CO2).</li>
</ol>
2. Staining of Reporter Cells with SNAP Substrate
<ol>
	<li>30 min prior to the beginning of the staining procedure allow aliquots of phenol red-free normal growth medium to warm up to 37 &deg;C.</li>
	<li>For easy handling of the SNAP substrate (in our case TMR Star) dissolve TMR Star in DMSO to obtain a concentration in the stock solution of 400 &mu;M, which can be stored at -20 &deg;C.</li>
	<li>Prior to staining, dilute 0.5 &mu;l of TMR Star stock solution in 200 &mu;l of phenol red-free normal growth medium (37 &deg;C) to obtain a final labeling concentration of 1 &mu;M.</li>
	<li>Remove normal growth medium from the asynchronously growing cells and incubate in labeling medium for 25 min under standard culture conditions.</li>
	<li>Remove labeling medium and wash cells four times with phenol red-free normal growth medium (37 &deg;C). Incubate cells in 300 &mu;l of phenol red-free normal growth medium (37 &deg;C) for 30 min. Prior to transporting to the microscope replace the medium with fresh phenol red-free normal growth medium (37 &deg;C) to remove residual unbound SNAP substrate.</li>
	<li>Transport cells to the microscope in a styrofoam box on a pre-warmed (37 &deg;C) heat block to minimize temperature variation.</li>
</ol>
3. Measurement of Fluorescence Intensity
<ol>
	<li>Two hours prior to the analysis adjust the air temperature of the climate chamber to 37 &deg;C in dry mode in order to bring the entire microscope with all its components to the desired temperature. Pre-heating before setting the humidity is important to avoid condensation and subsequent damage to the microscope.</li>
	<li>Adjust air humidity to 60% and CO2 to 5% prior to the start of the analysis.</li>
	<li>Start the Scan^R Acquisition software and define standard settings (see Table 1).</li>
	<li>Define the positions of the wells to be analyzed.</li>
	<li>Define the &Delta;t (acquisition cycle time) and the absolute number of acquisition cycles.</li>
	<li>If analysis of a higher number of wells is desired, select hardware autofocus, otherwise it is sufficient to use software autofocus alone.</li>
	<li>Start the acquisition and supervise for the first two acquisition cycles. The microscope will focus on the histone H2-GFP signal, with subsequent acquisition of a first image in that channel before the filter is changed and the corresponding TMR Star image is acquired (Figure 3). This is then repeated for all positions within a well and for each of the wells to be examined, before repeating again the next cycle.</li>
</ol>
4. Analysis of Proteolytic Profiles using Scan^R
<ol>
	<li>Start the Scan^R Analysis software and analyse the images with the cell nuclei, as visualized by histone H2-GFP, defined as the main object, using a threshold based on signal intensity and a watershed algorithm to assist in separating neighboring cells. A subobject consisting of a nucleus with cytoplasm should be created for TMRstar analysis. (Important properties of the main object are X and Y positions, time, and maximum and mean intensities of GFP for the main object and the total intensity divided by area for the TMRstar subobject.) This analysis process may take several hours due to large data quantities.</li>
	<li>Change to trace mode to visualize the subobject mean TMR Star fluorescence intensity over time (cell traces), assigned to the analyzed main objects (Figure 4). The number of cells to be examined can be narrowed down by gating on those measurements lasting at least 140 cycles and with a high maximum intensity of H2-GFP. Looking at larger numbers of cells allows a first representative and objective view (Figure 4).</li>
	<li>Select a cell trace of interest to visualize histone H2-GFP and TMR Star fluorescence simultaneously at the single cell level (Figure 5A).</li>
	<li>Using the right mouse click on a cell of interest and generate an exportable picture gallery for illustration of histone H2-GFP and cyclin B-SNAP TMR Star for every single time point.</li>
	<li>Change to the population mode of the Scan^R Analysis software and gate the region where the cell of interest is represented on the X vs. Y dot plot (Figure 5B).</li>
	<li>Apply a new dot plot window to the gated region and visualize mean TMR Star fluorescence intensity over time (Figure 5C).</li>
	<li>Export data (time and fluorescence intensity) to Microsoft Excel for further calculation.</li>
</ol>
5. Representative Results
Figure 5D and 5E depict cyclin B kinetics, represented by a TMR Star fluorescence intensity curve, of a cell that proceeds through a regular mitosis without signs of chromosomal misalignment (Figure 5E). Upon compaction of the cytoplasm following nuclear envelope breakdown (NEBD, as indicated by the red triangle), TMR Star fluorescence intensity shows an abrupt increase until the isomorphic window (brighter area in the diagram) is reached when the cell enters prometaphase 6. Fluorescence intensity remains at a stable level as long as the cell proceeds through prophase and metaphase and then starts to drop rapidly once all of the chromosomes have established a stable metaphase plate (Figure 5D and 5E). This drop precedes chromosome separation during anaphase (blue dot on the curve). In late mitosis the chromatin starts to decondense (blue bars) and the cell adopts interphase morphology while the fluorescence intensity curve approaches a plateau which is lower than the plateau before mitosis (Figure 5D and 5E).
<table border="1">
	<tbody>
		<tr>
			<td>Autofocus settings</td>
			<td>Histone H2-GFP (main object acquisition settings)</td>
			<td>Cyclin B-SNAP labeled with TMR Star 
			(acquisition settings)</td>
			<td>Acquisition cycle time Repetitions</td>
		</tr>
		<tr>
			<td>Coarse autofocus +/-39 &mu;m 24 Layers 
			Fine autofocus 
			+/-5.4 &mu;m 14 Layers</td>
			<td>GFP filter set: 
			Exposure time: 100 msec 
			Light intensity: 25%</td>
			<td>TRITC filter set: 
			Exposure time: 150 msec 
			Light intensity: 33.3%</td>
			<td>2 to 5 min.</td>
		</tr>
		<tr>
			<td>GFP filter set: 
			Exposure time: 12 msec 
			Light intensity: 12.5%</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Up to 48 hr of continued analysis (limited by reduced air humidity of 60%)</td>
		</tr>
	</tbody>
</table>
Table 1. Standard settings as used for analysis of cyclin B proteolysis. Histone H2-GFP was used as a reference structure to define the focus plane for measurement of cyclin B-SNAP fluorescence intensity. Light intensities during the focusing procedure are lower as compared to image acquisition settings to avoid cumulative phototoxicity.
<img alt="Figure 1" src="/files/ftp_upload/4239/4239fig1.jpg" /> 
 Figure 1. Schematic of the measurement of cyclin B proteolysis through expression of a chimeric cyclin B derivative with degradation characteristics similar to the endogenous protein. The decline in fluorescence intensity after pulse-chase labeling is a measure of APC/C-activity.
<img alt="Figure 2" src="/files/ftp_upload/4239/4239fig2.jpg" /> 
 Figure 2. Analysis of cells on 8 well slides or 96 well plates. Regular distribution of the cells across the surface of the well is achieved by resuspension in a final volume of 300 &mu;l and is recommended, if only a single or few positions are manually defined for analysis. Enrichment of cells in the center region is achieved by addition of cells to the center of prefilled or empty wells. This technique is recommended in case of automated image acquisition in different wells.
<img alt="Figure 3" src="/files/ftp_upload/4239/4239fig3.jpg" /> 
 Figure 3. Sequence of data acquisition. A) Detection of histone H2-GFP is used for focus plane definition and monitoring of chromosomal alignment during mitosis. B) Fluorescence intensity of the TMR Star fluorescence intensity is a measure for the absolute amount of pulse-chase labeled cyclin B-SNAP.
<img alt="Figure 4" src="/files/ftp_upload/4239/4239fig4.jpg" /> 
 Figure 4. Representative Scan^R Analysis software-based depiction of mean TMR Star fluorescence intensities (total TMRstar intensity divided by area) over time. Selection of a representative cell trace (blue). The corresponding images (shown on the right) allow the simultaneous visualization of histone H2-GFP (green) and cyclin B-SNAP (red). <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/4239/4239fig4large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 5" src="/files/ftp_upload/4239/4239fig5.jpg" /> 
 Figure 5. A+B) Example for gating of dots representing a cell of interest on the XY-dot plot. C) Visualization of the mean TMRstar fluorescence intensity (total TMRstar intensity divided by area) over time using the dot plot. D+E) Representative fluorescence intensity curve of cyclin B-SNAP with isomorphic window (brigher field on the diagram) between NEBD (nuclear envelope breakdown) and chromatin decondensation (blue bar) as generated in Microsoft Excel. Anaphase is indicated by the blue dot on the curve. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/4239/4239fig5large.jpg" target="_blank"> Click here to view larger figure</a>.
Movie 1. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/4239/4239movie1.avi" target="_blank"> Click here to view supplemental movie</a>.

<ol>
	<li>Musacchio, A., Salmon, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+spindle-assembly+checkpoint+in+space+and+time.">The spindle-assembly checkpoint in space and time.</a> Nat. Rev. Mol. Cell Biol. 8, 379-393 (2007).</li><li>Nasmyth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Segregating+sister+genomes:+the+molecular+biology+of+chromosome+separation.">Segregating sister genomes: the molecular biology of chromosome separation.</a> Science. 297, 559-565 (2002).</li><li>Cahill, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutations+of+mitotic+checkpoint+genes+in+human+cancers.">Mutations of mitotic checkpoint genes in human cancers.</a> Nature. 392, 300-303 (1998).</li><li>Lengauer, C., Kinzler, K. W., Vogelstein, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+instabilities+in+human+cancers.">Genetic instabilities in human cancers.</a> Nature. 396, 643-649 (1998).</li><li>Fletcher, D. A., Mullins, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+mechanics+and+the+cytoskeleton.">Cell mechanics and the cytoskeleton.</a> Nature. 463, 485-4892 (2010).</li><li>Schnerch, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+APC/C+activity+in+the+presence+of+chromosomal+misalignment+in+unperturbed+cell+populations.">Monitoring APC/C activity in the presence of chromosomal misalignment in unperturbed cell populations.</a> Cell Cycle. 11, (2012).</li><li>Keppler, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+general+method+for+the+covalent+labeling+of+fusion+proteins+with+small+molecules+in+vivo.">A general method for the covalent labeling of fusion proteins with small molecules in vivo.</a> Nat. Biotechnol. 21, 86-869 (2003).</li><li>Jansen, L. E., Black, B. E., Foltz, D. R., Cleveland, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Propagation+of+centromeric+chromatin+requires+exit+from+mitosis.">Propagation of centromeric chromatin requires exit from mitosis.</a> J. Cell. Biol. 176, 795-805 (2007).</li><li>Malureanu, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cdc20+hypomorphic+mice+fail+to+counteract+de+novo+synthesis+of+cyclin+B1+in+mitosis.">Cdc20 hypomorphic mice fail to counteract de novo synthesis of cyclin B1 in mitosis.</a> J. Cell Biol. 191, 313-329 (2010).</li><li>Mena, A. L., Lam, E. W., Chatterjee, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sustained+spindle-assembly+checkpoint+response+requires+de+novo+transcription+and+translation+of+cyclin+B1.">Sustained spindle-assembly checkpoint response requires de novo transcription and translation of cyclin B1.</a> PLoS One. 5, (2010).</li><li>Wolthuis, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cdc20+and+Cks+direct+the+spindle+checkpoint-independent+destruction+of+cyclin.">Cdc20 and Cks direct the spindle checkpoint-independent destruction of cyclin.</a> A. Mol. Cell. 30, 290-302 (2008).</li></ol>

No conflicts of interest declared.

We present here a live-cell imaging-based approach facilitating the simultaneous monitoring of cyclin B proteolysis and chromosome alignment. This approach allows the study of mitotic control in unperturbed cell populations at the single cell level. Cyclin B degradation curves facilitate direct insights into the activity of the APC/C and thus indirectly reflect the status of the SAC 6.
This approach, even though established based on the Scan^R software, can easily be reproduced on ...

<table><tbody><tr><td>Reporter cell line</td><td>generated in-house as described &lt;sup&gt;6&lt;/sup&gt;</td><td>clone 11 reporter cells&lt;br/&gt; (U2Os-based cyclin B-SNAP expressing cells)</td><td></td></tr><tr><td>Retroviral cyclin B-SNAP expression vector</td><td>generated in-house as described &lt;sup&gt;6&lt;/sup&gt;</td><td>pLNCX2-cyclin B mut5-SNAP</td><td></td></tr><tr><td>Phenolred-free DMEM</td><td>Gibco</td><td>21063-029</td><td>Supplementation with FCS, sodium pyruvate, penicillin/strepto-mycin required</td></tr><tr><td>SNAP-Cell TMR-Star</td><td>New England Biolabs</td><td>S9105S</td><td>Stock solution 400 &amp;mu;M in DMSO</td></tr><tr><td>Special Opstics Plate, 96 well</td><td>Costar</td><td>3720</td><td></td></tr><tr><td>&amp;mu;-Slide 8 well, ibiTreat</td><td>Ibidi</td><td>80826</td><td></td></tr><tr><td>Microscopy unit</td><td>Olympus</td><td>IX-81 inverse microscope with climate chamber</td><td></td></tr><tr><td>Objective</td><td>Olympus</td><td>UPLSAPO 20x objective (N.A. 0.75)</td><td></td></tr><tr><td>Acquisition software</td><td>Olympus</td><td>Scan^R Acquisition software (v.2.2.09)</td><td></td></tr><tr><td>Analysis software</td><td>Olympus</td><td>Scan^R Analysis software (v.1.2.0.6)</td><td></td></tr></tbody></table>

studying proteolysis of cyclin b at the single cell level in whole cell populations

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies during chromosome separation are a hallmark of malignancy and associated with progressive disease 2-4. The spindle assembly checkpoint (SAC) is a mitotic surveillance mechanism that holds back cells at metaphase until every single chromosome has established a stable bipolar attachment to the mitotic spindle1. The SAC exerts its function by interference with the activating APC/C subunit Cdc20 to block proteolysis of securin and cyclin B and thus chromosome separation and mitotic exit. Improper attachment of chromosomes prevents silencing of SAC signaling and causes continued inhibition of APC/CCdc20 until the problem is solved to avoid chromosome missegregation, aneuploidy and malignant growths1.
Most studies that addressed the influence of improper chromosomal attachment on APC/C-dependent proteolysis took advantage of spindle disruption using depolymerizing or microtubule-stabilizing drugs to interfere with chromosomal attachment to microtubules. Since interference with microtubule kinetics can affect the transport and localization of critical regulators, these procedures bear a risk of inducing artificial effects 5.
To study how the SAC interferes with APC/C-dependent proteolysis of cyclin B during mitosis in unperturbed cell populations, we established a histone H2-GFP-based system which allowed the simultaneous monitoring of metaphase alignment of mitotic chromosomes and proteolysis of cyclin B 6.
To depict proteolytic profiles, we generated a chimeric cyclin B reporter molecule with a C-terminal SNAP moiety 6 (Figure 1). In a self-labeling reaction, the SNAP-moiety is able to form covalent bonds with alkylguanine-carriers (SNAP substrate) 7,8 (Figure 1). SNAP substrate molecules are readily available and carry a broad spectrum of different fluorochromes. Chimeric cyclin B-SNAP molecules become labeled upon addition of the membrane-permeable SNAP substrate to the growth medium 7 (Figure 1). Following the labeling reaction, the cyclin B-SNAP fluorescence intensity drops in a pulse-chase reaction-like manner and fluorescence intensities reflect levels of cyclin B degradation 6 (Figure 1). Our system facilitates the monitoring of mitotic APC/C-dependent proteolysis in large numbers of cells (or several cell populations) in parallel. Thereby, the system may be a valuable tool to identify agents/small molecules that are able to interfere with proteolytic activity at the metaphase to anaphase transition. Moreover, as synthesis of cyclin B during mitosis has recently been suggested as an important mechanism in fostering a mitotic block in mice and humans by keeping cyclin B expression levels stable 9,10, this system enabled us to analyze cyclin B proteolysis as one element of a balanced equilibrium 6.

Watch this Scientific Journal Video about Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations at JoVE.com

Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations

Metaphase to anaphase transition is triggered through anaphase-promoting complex (APC/C)-dependent ubiquitination and subsequent destruction of cyclin B. Here, we established a system which, following pulse-chase labeling, allows monitoring cyclin B proteolysis in entire cell populations and facilitates the detection of interference by the mitotic checkpoint.

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies ...

studying-proteolysis-cyclin-b-at-single-cell-level-whole-cell

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

10.4K Views.  University Medical Center Freiburg. Metaphase to anaphase transition is triggered through anaphase-promoting complex (APC/C)-dependent ubiquitination and subsequent destruction of cyclin B. Here, we established a system which, following pulse-chase labeling, allows monitoring cyclin B proteolysis in entire cell populations and facilitates the detection of interference by the mitotic checkpoint.

Video: Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations

Single-cell Gene Expression Profiling Using FACS and qPCR with Internal Standards

Gene expression measurements from bulk populations of cells can obscure the considerable transcriptomic variation of individual cells within those populations. Single-cell gene expression measurements can help assess the role of noise in gene expression, identify correlations in the expression of pairs of genes, and reveal subpopulations of cells that respond differently to a stimulus. Here, we describe a procedure to measure the expression of up to 96 genes in single mammalian cells isolated from a population growing in tissue culture. Cells are sorted into lysis buffer by fluorescence-activated cell sorting (FACS), and the mRNA species of interest are reverse-transcribed and amplified. Gene expression is then measured using a microfluidic real-time PCR machine, which performs up to 96 qPCR assays on up to 96 samples at a time. We also describe the generation and use of PCR amplicon standards to enable the estimation of the absolute number of each transcript. Compared with other methods of measuring gene expression in single cells, this approach allows for the quantification of more distinct transcripts than RNA FISH at a lower cost than RNA-Seq.

We describe a method to sort single mammalian cells and to quantify the expression of up to 96 target genes of interest in each cell. This method includes the use of internal qPCR standards to enable the estimation of absolute transcript counts.

Gene expression measurements from bulk populations of cells can obscure the considerable transcriptomic variation of individual cells within those ...

Genetics

Identification of Cyclin-dependent Kinase 1 Specific Phosphorylation Sites by an In Vitro Kinase Assay

Cyclin-dependent kinase 1 (Cdk1) is a master controller for the cell cycle in all eukaryotes and phosphorylates an estimated 8 - 13% of the proteome; however, the number of identified targets for Cdk1, particularly in human cells is still low. The identification of Cdk1-specific phosphorylation sites is important, as they provide mechanistic insights into how Cdk1 controls the cell cycle. Cell cycle regulation is critical for faithful chromosome segregation, and defects in this complicated process lead to chromosomal aberrations and cancer.
Here, we describe an in vitro kinase assay that is used to identify Cdk1-specific phosphorylation sites. In this assay, a purified protein is phosphorylated in vitro by commercially available human Cdk1/cyclin B. Successful phosphorylation is confirmed by SDS-PAGE, and phosphorylation sites are subsequently identified by mass spectrometry. We also describe purification protocols that yield highly pure and homogeneous protein preparations suitable for the kinase assay, and a binding assay for the functional verification of the identified phosphorylation sites, which probes the interaction between a classical nuclear localization signal (cNLS) and its nuclear transport receptor karyopherin &#945;. To aid with experimental design, we review approaches for the prediction of Cdk1-specific phosphorylation sites from protein sequences. Together these protocols present a very powerful approach that yields Cdk1-specific phosphorylation sites and enables mechanistic studies into how Cdk1 controls the cell cycle. Since this method relies on purified proteins, it can be applied to any model organism and yields reliable results, especially when combined with cell functional studies.

Identification of Cyclin-dependent Kinase 1 Specific Phosphorylation Sites by an In Vitro Kinase Assay

Cyclin-dependent kinase 1 (Cdk1) is activated in the G2 phase of the cell cycle and regulates many cellular pathways. Here, we present a protocol for an in vitro kinase assay with Cdk1, which allows the identification of Cdk1-specific phosphorylation sites for establishing cellular targets of this important kinase.

Cyclin-dependent kinase 1 (Cdk1) is a master controller for the cell cycle in all eukaryotes and phosphorylates an estimated 8 - 13% of the proteome; ...

Biochemistry

Reconstitution of Cell-cycle Oscillations in Microemulsions of Cell-free Xenopus Egg Extracts

Real-time measurement of oscillations at the single-cell level is important to uncover the mechanisms of biological clocks. Although bulk extracts prepared from Xenopus laevis eggs have been powerful in dissecting biochemical networks underlying the cell-cycle progression, their ensemble average measurement typically leads to a damped oscillation, despite each individual oscillator being sustained. This is due to the difficulty of perfect synchronization among individual oscillators in noisy biological systems. To retrieve the single-cell dynamics of the oscillator, we developed a droplet-based artificial cell system that can reconstitute mitotic cycles in cell-like compartments encapsulating cycling cytoplasmic extracts of Xenopus laevis eggs. These simple cytoplasmic-only cells exhibit sustained oscillations for over 30 cycles. To build more complicated cells with nuclei, we added demembranated sperm chromatin to trigger nuclei self-assembly in the system. We observed a periodic progression of chromosome condensation/decondensation and nuclei envelop breakdown/reformation, like in real cells. This indicates that the mitotic oscillator functions faithfully to drive multiple downstream mitotic events. We simultaneously tracked the dynamics of the mitotic oscillator and downstream processes in individual droplets using multi-channel time-lapse fluorescence microscopy. The artificial cell-cycle system provides a high-throughput framework for quantitative manipulation and analysis of mitotic oscillations with single-cell resolution, which likely provides important insights into the regulatory machinery and functions of the clock.

Reconstitution of Cell-cycle Oscillations in Microemulsions of Cell-free Xenopus Egg Extracts

We present a method for the generation of in vitro self-sustained mitotic oscillations at the single-cell level by encapsulating egg extracts of Xenopus laevis in water-in-oil microemulsions.

Real-time measurement of oscillations at the single-cell level is important to uncover the mechanisms of biological clocks. Although bulk extracts ...

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.

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 ...

Developmental Biology

Ordering Single Cells and Single Embryos in 3D Confinement: A New Device for High Content Screening

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening based on cells in such environments have therefore serious limitations: cell organelles show extreme phenotypes, cell morphologies and sizes are heterogeneous and/or specific cell organelles cannot be properly visualized. In addition, cells in vivo are located in a 3D environment; in this situation, cells show different phenotypes mainly because of their interaction with the surrounding extracellular matrix of the tissue. In order to standardize and generate order of single cells in a physiologically-relevant 3D environment for cell-based assays, we report here the microfabrication and applications of a device for in vitro 3D cell culture. This device consists of a 2D array of microcavities (typically 105 cavities/cm2), each filled with single cells or embryos. Cell position, shape, polarity and internal cell organization become then normalized showing a 3D architecture. We used replica molding to pattern an array of microcavities, &#8216;eggcups&#8217;, onto a thin polydimethylsiloxane (PDMS) layer adhered on a coverslip. Cavities were covered with fibronectin to facilitate adhesion. Cells were inserted by centrifugation. Filling percentage was optimized for each system allowing up to 80%. Cells and embryos viability was confirmed. We applied this methodology for the visualization of cellular organelles, such as nucleus and Golgi apparatus, and to study active processes, such as the closure of the cytokinetic ring during cell mitosis. This device allowed the identification of new features, such as periodic accumulations and inhomogeneities of myosin and actin during the cytokinetic ring closure and compacted phenotypes for Golgi and nucleus alignment. We characterized the method for mammalian cells, fission yeast, budding yeast, C. elegans with specific adaptation in each case. Finally, the characteristics of this device make it particularly interesting for drug screening assays and personalized medicine.

We report a device and a new method to study cells and embryos. Single cells are precisely ordered in microcavity arrays. Their 3D confinement is a step towards 3D environments encountered in physiological conditions and allows organelle orientation. By controlling cell shape, this setup minimizes variability reported in standard assays.

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening ...

Bioengineering

Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such probes typically consist of variants of CFP and YFP, intervened by a phosphorylatable sequence and a phospho-binding domain. Upon phosphorylation, the probe changes conformation, which results in a change of the distance or orientation between CFP and YFP, leading to a change in FRET efficiency (Fig 1). Several probes have been published during the last decade, monitoring the activity balance of multiple kinases and phosphatases, including reporters of PKA2, PKB3, PKC4, PKD5, ERK6, JNK7, Cdk18, Aurora B9 and Plk19. Given the modular design, additional probes are likely to emerge in the near future10. 
Progression through the cell cycle is affected by stress signaling pathways 11. Notably, the cell cycle is regulated differently during unperturbed growth compared to when cells are recovering from stress12.Time-lapse imaging of cells through the cell cycle therefore requires particular caution. This becomes a problem particularly when employing ratiometric imaging, since two images with a high signal to noise ratio are required to correctly interpret the results. Ratiometric FRET imaging of cell cycle dependent changes in kinase and phosphatase activities has predominately been restricted to sub-sections of the cell cycle8,9,13,14.
Here, we discuss a method to monitor FRET-based probes using ratiometric imaging throughout the human cell cycle. The method relies on equipment that is available to many researchers in life sciences and does not require expert knowledge of microscopy or image processing.

FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such ...

Analysis of Cell Cycle Position in Mammalian Cells

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of cell proliferation underlies the pathology of diseases like cancer. As such there is great need to be able to investigate cell proliferation and quantitate the proportion of cells in each phase of the cell cycle. It is also of vital importance to indistinguishably identify cells that are replicating their DNA within a larger population. Since a cell&prime;s decision to proliferate is made in the G1 phase immediately before initiating DNA synthesis and progressing through the rest of the cell cycle, detection of DNA synthesis at this stage allows for an unambiguous determination of the status of growth regulation in cell culture experiments.
DNA content in cells can be readily quantitated by flow cytometry of cells stained with propidium iodide, a fluorescent DNA intercalating dye. Similarly, active DNA synthesis can be quantitated by culturing cells in the presence of radioactive thymidine, harvesting the cells, and measuring the incorporation of radioactivity into an acid insoluble fraction. We have considerable expertise with cell cycle analysis and recommend a different approach. We Investigate cell proliferation using bromodeoxyuridine/fluorodeoxyuridine (abbreviated simply as BrdU) staining that detects the incorporation of these thymine analogs into recently synthesized DNA. Labeling and staining cells with BrdU, combined with total DNA staining by propidium iodide and analysis by flow cytometry1 offers the most accurate measure of cells in the various stages of the cell cycle. It is our preferred method because it combines the detection of active DNA synthesis, through antibody based staining of BrdU, with total DNA content from propidium iodide. This allows for the clear separation of cells in G1 from early S phase, or late S phase from G2/M. Furthermore, this approach can be utilized to investigate the effects of many different cell stimuli and pharmacologic agents on the regulation of progression through these different cell cycle phases.
In this report we describe methods for labeling and staining cultured cells, as well as their analysis by flow cytometry. We also include experimental examples of how this method can be used to measure the effects of growth inhibiting signals from cytokines such as TGF-&beta;1, and proliferative inhibitors such as the cyclin dependent kinase inhibitor, p27KIP1. We also include an alternate protocol that allows for the analysis of cell cycle position in a sub-population of cells within a larger culture5. In this case, we demonstrate how to detect a cell cycle arrest in cells transfected with the retinoblastoma gene even when greatly outnumbered by untransfected cells in the same culture. These examples illustrate the many ways that DNA staining and flow cytometry can be utilized and adapted to investigate fundamental questions of mammalian cell cycle control.

Determining the cell cycle position of a population of cells, or understanding how signals affect proliferation, can be readily measured by flow cytometry using this protocol. We report a simple experimental approach to staining cells and quantifying their position in the cell cycle.

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The nuclear-encoded proteins are transported into mitochondria guided by their mitochondria targeting sequences (MTS); however, a majority of mitochondrial localized proteins lack an identifiable MTS. Nevertheless, the fact that MTS can instruct proteins to go into the mitochondria provides a valuable tool for studying mitochondrial functions of normally nuclear and/or cytoplasmic proteins. We have recently identified the cell cycle kinase CyclinB1/Cdk1 complex in the mitochondria. To specifically study the mitochondrial functions of this complex, mitochondrial overexpression and knock-down of this complex without interfering with its nuclear or cytoplasmic functions were essential. By tagging CyclinB1/Cdk1 with MTS, we were able to achieve mitochondrial overexpression of this complex to study its mitochondrial targets as well as functions. Via tagging dominant-negative Cdk1 with MTS, inhibition of Cdk1 activity was accomplished particularly in the mitochondria. Potential mitochondrial targets of CyclinB1/Cdk1 complex were identified using a gel-based proteomics approach. Unlike traditional 2D gel analysis, we employed 2-dimensional difference gel electrophoresis (2D-DIGE) technology followed by phosphoprotein staining to fluorescently label differentially phosphorylated proteins in mitochondrial Cdk1 expressing cells. Identification of phosphoprotein spots that were altered in wild type versus dominant negative Cdk1 bearing mitochondria revealed the identity of mitochondrial targets of Cdk1. Finally, to determine the effect of CyclinB1/Cdk1 mitochondrial localization in cell cycle progression, a cell proliferation assay using a synthetic thymidine analogue EdU (5-ethynyl-2&#8242;-deoxyuridine) was used to monitor the cells as they go through the cell cycle and replicate their DNA. Altogether, we demonstrated a variety of approaches available to study mitochondrial localization and activity of a cell cycle kinase. These are advanced, yet easy to follow methods that will be beneficial to many cell biology researchers.

Here, we outline how to study mitochondrial localization of a (cell cycle) kinase, and how to determine its sub-mitochondrial location as well as potential mitochondrial substrates/targets. Forced expression of proteins into the mitochondria provides a useful tool for studying the functional consequences of mitochondrial localization of a protein of interest.

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The ...

Cycloheximide Chase Analysis of Protein Degradation in Saccharomyces cerevisiae

Regulation of protein abundance is crucial to virtually every cellular process. Protein abundance reflects the integration of the rates of protein synthesis and protein degradation. Many assays reporting on protein abundance (e.g., single-time point western blotting, flow cytometry, fluorescence microscopy, or growth-based reporter assays) do not allow discrimination of the relative effects of translation and proteolysis on protein levels. This article describes the use of cycloheximide chase followed by western blotting to specifically analyze protein degradation in the model unicellular eukaryote, Saccharomyces cerevisiae (budding yeast). In this procedure, yeast cells are incubated in the presence of the translational inhibitor cycloheximide. Aliquots of cells are collected immediately after and at specific time points following addition of cycloheximide. Cells are lysed, and the lysates are separated by polyacrylamide gel electrophoresis for western blot analysis of protein abundance at each time point. The cycloheximide chase procedure permits visualization of the degradation kinetics of the steady state population of a variety of cellular proteins. The procedure may be used to investigate the genetic requirements for and environmental influences on protein degradation.

Cycloheximide Chase Analysis of Protein Degradation in Saccharomyces cerevisiae

Protein abundance reflects the rates of both protein synthesis and protein degradation. This article describes the use of cycloheximide chase followed by western blotting to analyze protein degradation in the model unicellular eukaryote, Saccharomyces cerevisiae (budding yeast).

Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations

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Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

Analysis of Cell Cycle Position in Mammalian Cells

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1

Cycloheximide Chase Analysis of Protein Degradation in Saccharomyces cerevisiae