YS-T is supported by the European Research Council (ERC), the Israel Science Foundation (ISF) (250/06), ISF-Bikura, Israel Cancer Research Fund (ICRF), German-Israeli Foundation for Scientific Research and Development (GIF), USA-Israel Binational Science Foundation (BSF), German-Israeli Project Cooperation (DIP), and the Israeli Ministries of Science and Health, and is the Jane Stern Lebell Family Fellow in Life Sciences at Bar-Ilan University. YB is grateful to the Azrieli Foundation for the award of an Azrieli fellowship.

The cell system used must contain an integrated gene construct that includes MS2 repeat sequences for tagging the mRNA in living cells. In this protocol we use a human U2OS cell line harboring a stably integrated &beta;-actin gene, that is fluorescently labeled at the DNA, RNA and protein levels8, as follows (Figure 1A): The 5&#x2019; of the gene contains lac operator (lacO) repeats &#x2013; these are used to tag the gene (DNA) and thereby identify the genomic locus of integration. A fluorescently tagged lac repressor protein (LacI) will bind to the lacO repeats and tag the DNA. The mRNAs are tagged with YFP-MS2 proteins that bind to the MS2 repeats (stem-loops). The gene product is CFP-actin and therefore induction of the gene will result in the appearance of CFP-tagged actin cytoskeleton. The gene construct is transfected into a U2OS Tet-On stable cell line and its expression is under Tet transcriptional control.
1. Cell plating and transfection:
<ol>
 <li>48 hours before the measurements, seed &tilde;2*105 cells on glass bottomed tissue culture dishes (35 mm petri dishes with a 14 mm glass-bottomed microwell (0.16-0.19 mm thickness; Cat. No. P35G-1.5-14-C, MatTek, Ashland, MA)). Add 2ml of medium to the cells, in order to reach 50&#x2013;80% confluence.</li>
 <li>24 hours before the experiment, transiently transfect the cells with constructs that express the fluorescent markers for tagging the DNA and the mRNA. Use FuGENE6 transfection reagent (Roche). 
 The constructs used:<ol type="a"> 
 <li>YFP-MS2-NLS will tag the MS2 stem-loops in the mRNA transcripts. The nuclear localization sequence (NLS) provides the nuclear entry sequence for the YFP-MS2 protein.</li>
 <li>optional - CFP-LacI or RFP-LacI that will bind the lacO repeats in the 5&#x2019; of the gene and will label the gene locus. Tagging the gene locus assists in tracking the gene during the FRAP experiment but is not necessary.</li></ol></li>
 <li>Incubate the cells for at least 12 hrs, to obtain good expression of the transient transfected proteins (see note).</li>
 <li>6 - 12 hours before the experiment add 1.5 &mu;g/ml doxycycline (Sigma) to the cells for activating of the Tet-inducible CFP-actin gene.</li> 
</ol>
Notes: 
* It is important not to reach very high expression levels of the transfected proteins so as not to mask the signal obtained specifically from the transcription site, relative to the background signal. Levels of expression can be controlled by: a) adjusting the time between transfection and the experiment; b) changing the strength of the promotor (e.g. CMV is stronger than the L30 promoter); c) adding a nuclear export signal (NES) to the YFP-MS2-NLS protein, which will help distribute the YFP signal in the whole cell.

* The fluorophore used for the measurements should preferably be photostable throughout long acquisitions; therefore green/yellow emitting fluorophores are better than red fluorophores. Information on the different fluorophores can be found in ref 10. 
* Determine that the different light channels do not bleed-through to each other. Use slides containing specimens labeled with each of the markers separately, measure in each slide the intensity of both channels, and then calculate the bleed-through.
2. FRAP:
The experiment can be performed on any microscope that is capable of both acquiring images and performing photobleaching with a laser beam. Typically, FRAP experiments are performed with a confocal laser scanning microscope (CLSM). However, since in our case the experiment is relatively long (15-30 min), and we are following one small locus within the nucleus, it is important to be able to keep the gene locus/transcription site in focus throughout the whole experiment. This requires rapid imaging in 3 dimensions over time (4D), and such acquisition rates are hard to obtain using a scanning confocal microscope. Therefore, we use a 3D-FRAP system (Photometrics) built on a wide-field Olympus IX81 microscope (63X Plan-Apo, 1.4 NA) equipped with an EM-CCD (Quant-EM, Roper) that can acquire images at high speed (up to 30 images/sec). The system has 405 nm and 491 nm lasers that can be synchronized with the camera to enable photobleaching on a specific region of interest (ROI) of the sample. The microscope is equipped with a Lambda DG-4 light source (Sutter) and a XY&amp;Z stage (Prior) that allows fast and accurate imaging in the Z-axis. All the equipment is integrated and controlled by MetaMorph (Molecular Devices). The live-cell experiments are performed at 37&deg;C with 5% CO2 using a live-cell chamber system (Tokai).
<ol>
 <li>Procedure: Half an hour before the experiment, turn on the heating device, microscope and laser. During the experiment the temperature needs to remain constant, otherwise it might cause focus changes during the long acquisition times. Laser power must be stable for consistent power output.</li>
 <li>Synchronize the laser with the camera:
 <ol>
 <li>Search for an area in the plate that does not contain cells (using DIC).</li>
 <li>Use the same light channel that will be used for the experiments. Here we use the 491 nm laser for the YFP channel.</li>
 <li>Point the laser to the middle of the image.</li>
 <li>Use the synchronize button (provided macro).</li>
 <li>Try different spots in the field to ensure that the laser and camera are synchronized.</li>
 </ol>
 </li>
 <li>Choose a good candidate cell for the experiment. Although we work with stable cell lines, we find that not all cells have similar expression levels. Good candidates would be cells that: a) exhibit a clear YFP-MS2-tagged transcription site above the diffuse YFP-MS2 background; and b) have enough free YFP-MS2 protein in the cell so as to be able to recover completely after the photobleaching (example in figure 1B - compare the two cells; the left cell is not a good candidate for bleaching).</li> 
 <li>Choose appropriate measuring conditions. The FRAP procedure requires the acquiring of many images, and therefore it is important to balance between reasonable exposure times and using as low light as possible. High light levels might affect the cell due to phototoxicity, and will cause photobleaching. It is recommended to keep imaging parameters constant between different experiments. 
 Cells are imaged in the YFP channel with 150 msec exposure times, and in the CFP channel for the detection of CFP-lacI (genomic locus). Following the CFP-LacI tagged locus allows the detection of the gene even when the YFP-MS2 mRNA signal is photobleached (optional). For each acquisition, 7 Z-slices are taken every 350 nm. The active transcription site is bleached using the 491 nm laser.</li> 
 <li>Perform the full experiment but without bleaching. The FRAP experiment measures the time it takes for the transcription site to return to steady state. First, we want to verify that the active transcription site is functional and stable at steady state. Therefore, we perform a complete FRAP experiment but without bleaching to ensure that the measured intensity of the transcription site does not change during the experiment time frame. We use this experiment to also test our imaging conditions by measuring the total photobleaching caused by the full imaging series. As a rule of thumb, the full movie photobleach should not be more than 30% of the initial intensity. This can be measured by using the intensity ratio of the same spot in the nucleus from the last image, divided by first image (ratio should be larger than 0.7).</li>
 <li>FRAP: The YFP-MS2 signal at the active transcription site is bleached using the 491 nm laser. 6 pre-bleach images are acquired. Post-bleach images are acquired in a sequence of 3 time frequencies: 15 images every 3 sec, 15 images every 6 sec, and 26 (or 45 for long experiments) images every 30 sec. The changes in frequency of acquiring images allows to acquire more images during the times that show the highest intensity change (initial recovery), and less images towards the end when the changes in signal are less prominent.</li> 
</ol>
Notes: This experiment differs from "regular" FRAP in that: 
* FRAP of mRNA at the transcription site measures the binding of YFP-MS2 proteins to the nascent mRNA, whereas typical FRAP experiments measure the mobility of different proteins in the cell. Therefore, there is no interest in measuring the diffusive pool of YFP-MS2 proteins. When the time between bleaching and the first acquired image is relatively long it is possible to disregard this diffusing pool of free YFP-MS2 proteins.
* Since the recovery of the YFP-MS2 signal is relatively long (15-30 min), it is important to keep the transcription site in focus. That is why a series of Z-slices is acquired at every time point.
3. Normalization of the FRAP data:
The 4D FRAP data is analyzed using the free Image analysis software ImageJ (NIH, Bethesda, MD; <a href="http://rsb.info.nih.gov/ij/">http://rsb.info.nih.gov/ij/</a>). Following is analysis performed with macros available through the ImageJ software, but of course self-written macros can be generated.
<ol>
 <li>Z-stacks are maximum projected at every time-point, resulting in a 2D movie over time for further analysis.</li> 
 <li>The transcription site is tracked in each frame and the mean intensity is measured using the Spot Tracker tool.11 The data is saved in an Excel sheet for further analysis.</li>
 <li>For every time-point, the mean intensities at other regions of interest (ROI) are measured as follows (using the Time Series Analyzer plug-in of ImageJ): a) The background is taken from an ROI outside of the cell = B(t). b) An ROI in the nucleoplasm, but as far as possible from the bleach site = C(t). c) The transcription site = S(t) (as measured by the spot tracker).</li> 
 <li>Correct the data for photobleaching caused during imaging: Subtract the background from all other measurements and then correct the bleach. Sc(t) is the corrected intensity of the transcription site at time t.</li> 
</ol>
 eq. 1: <img src="/files/ftp_upload/2898/2898eq1.jpg" alt="Equation 1" />
<ol start="5">
<li>Normalize the data: first calculate the average of the pre-bleach images, which shows the intensity of the transcription site at steady-state ( = &lt;Sc(pre-bleach)&gt;). </li>
</ol>
Then normalize the data:
eq. 2 <img src="/files/ftp_upload/2898/2898eq2.jpg" alt="Equation 2" />
 The data are normalized for at least 10 experiments. Calculate the average of the experiments and the standard deviation (STD) for the error-bars at every time-point (Figure 1C). 
Notes: 
The diffusion of the YFP-MS2 protein during the FRAP analysis was disregarded since the diffusion rate of free nucleoplasmic YFP-MS2 is very rapid, while the bound YFP-MS2 is associated with high affinity to the mRNA, and does not reattach to the transcription site. This is why the first image after bleaching should be removed before analysis. 
4. Analyzing the data using a mathematical model: 
In order to retrieve the kinetic parameters from this kind of data set, we used the simplest described model that can fit this kind of data. The full explanation of this specific model can be read here5. The model contains the following equations that describe the scheme in figure 2A:
<img src="/files/ftp_upload/2898/2898eq3.jpg" alt="Equation 3" /> 
<img src="/files/ftp_upload/2898/2898eq4.jpg" alt="Equation 4" /> 
Having the model parameters, it is possible to calculate the elongation rate:
<img src="/files/ftp_upload/2898/2898eq5.jpg" alt="Equation 5" /> 
We analyzed the curve using Berkeley Madonna (<a href="http://www.berkeleymadonna.com">http://www.berkeleymadonna.com</a>), as previously modeled5 but other options are available (MatLab, Virtual Cell, COPASI etc.). The data was fit to the model and rate constants were extracted. We attach the code in Berkeley Madonna. In order to best fit the data we calculated the steady state and then fit the normalized experimental data to the normalized model. 

5. Representative Results:
We photobleached the labeled mRNA signal at the active transcription site using high laser power, and followed the recovery of the YFP-MS2- signal over time. The fluorescent signal at the transcription site consists of the steady state kinetics of mRNA synthesis as well as mRNA released from the site. Measuring transcription site intensity over time without bleaching gives a 'constant' signal (figure 1C, the red dots) which implies that this gene is currently in a relative steady state. This means that the signal that accumulates on the newly transcribed mRNA is equal to the signal that leaves when an mRNA is released from the transcription site. When photobleached, this system continues in its steady-state, but is now possible to measure the time it takes for the signal to recover. Therefore, the recovery of fluorescence mostly depends on the elongation properties of the polymerase. The 3D FRAP system allowed the acquisition of the full 3D nuclear volume during FRAP recovery, and thereby no signal was lost during the experiments. For calculating the kinetic parameters from the data, the normalized curve was fitted to the kinetic model describing elongation, that is based on differential equations.5 The advantage of using a simple ODE model is that it can be fit to only a small number of parameters. The initial step of the model (entry point) is elongation, namely the process responsible for synthesizing new MS2-binding sites. The end step (exit point) is mRNA release into the nucleoplasm. The model is based on the recovery curves, which consist of two kinetically resolved components, one fast and one slower. The fast component refers to elongation, whereas the slower component was modeled as polymerase pausing during elongation. The two polymerase states were modeled as elongation with a stochastic transition to pausing. We optimized the differential equations from this model, which yielded an elongation speed of 3.3 kb/minute with a stochastic transition to a slower synthesis rate (pausing for a cumulative time of 2.5 minutes). Modeling showed that this transition from elongation to pausing affected only 11% of the polymerases that entered elongation. 
<table border="1">
 <tbody>
 <tr>
 <td>Kout</td>
 <td>0.0112</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>K pause_in</td>
 <td>0.0015</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>K pause_out</td>
 <td>0.0067</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Elongation time</td>
 <td>(Kout + k pause_out)-1</td>
 <td>56 sec</td>
 </tr>
 <tr>
 <td>Pause time </td>
 <td>(k pause_out)-1</td>
 <td>2.5 minutes</td>
 </tr>
 <tr>
 <td>Pause probability</td>
 <td>(K pause_in) / (K pause_in + kout)</td>
 <td>11 %</td>
 </tr>
 </tbody>
</table> 
Table 1.The model results in a parameter table.
<img src="/files/ftp_upload/2898/2898fig1.jpg" alt="Figure 1" /> 
Figure 1. (A) The experimental cell system used for following gene expression in vivo. Schematic representation of the CFP-actin gene 
construct. The 5'-end contains a series of 256 lacO repeats that are bound by RFP-LacI (red) and mark the site of gene integration. Transcriptional 
induction from the minimal CMV promoter is achieved by the binding of reverse tetracycline transcriptional activator (rtTA or Tet-On) to Tet-responsive elements 
(TREs) in the presence of dox. The transcribed mRNA contains the coding sequence for CFP-&beta;-actin, and 24 MS2 repeats that are bound by the YFP-MS2 fusion 
protein. At the 3'-end, a portion of a rabbit &beta;-globin exon-intron-exon module is followed by a cleavage-polyA signal. The scheme also describes the elongation 
assay showing fluorescent MS2 proteins (yellow) attached to the newly transcribed mRNAs transcribed by RNA Pol II. (B) An active transcription 
site (YFP-MS2, right hand cell) was photobleached and the fluorescence recovery was tracked over time (frames on the bottom). We measured the fluorescence 
intensity of both bleach (middle line) and non-bleached (bottom line) active transcription sites. Note the left-hand cell is not suitable for a photobleaching 
experiment. (C) The experimental measurements performed on the CFP-actin gene transcription sites. In blue the recovery curves of the YFP-MS2 
FRAP measurements. In red, a non-bleached cell from the same experiment showing the YFP-MS2 signal at the transcription site at steady state.
<img src="/files/ftp_upload/2898/2898fig2.jpg" alt="Figure 2" /> 
Figure 2. (A) Scheme of the model describing the entrance and exit kinetics of mRNAs marked with YFP-MS2 molecules at the transcription 
site. (B) YFP-MS2 FRAP data and the fitted simulation (the average of ten experiments was fitted to the model). The calculated residuals of the fit 
are presented at the bottom.

<ol>
	<li>Wada, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+wave+of+nascent+transcription+on+activated+human+genes.">A wave of nascent transcription on activated human genes.</a> Proc. Natl. Acad. Sci. U. S. A. 106, 18357-18361 (2009).</li><li>Singh, J., Padgett, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rates+of+in+situ+transcription+and+splicing+in+large+human+genes.">Rates of in situ transcription and splicing in large human genes.</a> Nat Struct Mol Biol. 16, 1128-1133 (2009).</li><li>Brody, Y., Shav-Tal, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+transcription+in+real-time.">Visualizing transcription in real-time.</a> Cent Eur J Biol. 3, 11-18 (2008).</li><li>Lamond, A. 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No conflicts of interest declared.

The method for measuring polymerase kinetic activity in living cells can be divided into two parts. The first part describes the 'wet' procedure which enables the measuring and retrieving of the kinetic data of mRNA transcription, whereas in the second part, the data is analyzed using an ODE model.5 A number of studies have now utilized this approach for extracting transcription kinetics in living cells5, 6, 8, 12-14. The major difference between these studies was the way the FRAP recovery curve was...

<table border="1">
	<tbody>
		<tr>
			<th>Name of the reagent </th>
			<th>Company </th>
		</tr>
		<tr>
			<td>Doxycycline</td>
			<td>Sigma</td>
		</tr>
		<tr>
			<td>FuGENE 6 transfection reagent</td>
			<td>Roche</td>
		</tr>
	</tbody>
</table>

measuring the kinetics of mrna transcription in single living cells

The transcriptional activity of RNA polymerase II (Pol II) is a dynamic process and therefore measuring the kinetics of the transcriptional process in vivo is of importance. Pol II kinetics have been measured using biochemical or molecular methods.1-3 In recent years, with the development of new visualization methods, it has become possible to follow transcription as it occurs in real time in single living cells.4 Herein we describe how to perform analysis of Pol II elongation kinetics on a specific gene in living cells.5, 6 Using a cell line in which a specific gene locus (DNA), its mRNA product, and the final protein product can be fluorescently labeled and visualized in vivo, it is possible to detect the actual transcription of mRNAs on the gene of interest.7, 8 The mRNA is fluorescently tagged using the MS2 system for tagging mRNAs in vivo, where the 3'UTR of the mRNA transcripts contain 24 MS2 stem-loop repeats, which provide highly specific binding sites for the YFP-MS2 coat protein that labels the mRNA as it is transcribed.9 To monitor the kinetics of transcription we use the Fluorescence Recovery After Photobleaching (FRAP) method. By photobleaching the YFP-MS2-tagged nascent transcripts at the site of transcription and then following the recovery of this signal over time, we obtain the synthesis rate of the newly made mRNAs.5 In other words, YFP-MS2 fluorescence recovery reflects the generation of new MS2 stem-loops in the nascent transcripts and their binding by fluorescent free YFP-MS2 molecules entering from the surrounding nucleoplasm. The FRAP recovery curves are then analyzed using mathematical mechanistic models formalized by a series of differential equations, in order to retrieve the kinetic time parameters of transcription.

Watch this Scientific Journal Video about Measuring the Kinetics of mRNA Transcription in Single Living Cells at JoVE.com

Measuring the Kinetics of mRNA Transcription in Single Living Cells

RNA polymerase II transcriptional kinetics are measured on specific genes in living cells. mRNAs transcribed from the gene of interest are fluorescently tagged and using Fluorescence Recovery After Photobleaching (FRAP) the in vivo kinetics of transcriptional elongation are obtained.

The transcriptional activity of RNA polymerase II (Pol II) is a dynamic process and therefore measuring the kinetics of the transcriptional process in ...

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15.4K Views.  Bar-Ilan University. RNA polymerase II transcriptional kinetics are measured on specific genes in living cells. mRNAs transcribed from the gene of interest are fluorescently tagged and using Fluorescence Recovery After Photobleaching (FRAP) the in vivo kinetics of transcriptional elongation are obtained.

Video: Measuring the Kinetics of mRNA Transcription in Single Living Cells

Method for Measurement of Viral Fusion Kinetics at the Single Particle Level

Membrane fusion is an essential step during entry of enveloped viruses into cells. Conventional fusion assays typically report on a large number of fusion events, making it difficult to quantitatively analyze the sequence of the molecular steps involved. We have developed an in vitro, two-color fluorescence assay to monitor kinetics of single virus particles fusing with a target bilayer on an essentially fluid support.
Influenza viral particles are incubated with a green lipophilic fluorophore to stain the membrane and a red hydrophilic fluorophore to stain the viral interior. We deposit a ganglioside-containing lipid bilayer on the dextran-functionilized glass surface of a flow cell, incubate the viral particles on the planar bilayer and image the fluorescence of a 100 x 100 &mu;m2 area, containing several hundreds of particles, on a CCD camera. By imaging both the red and green fluorescence, we can simultaneously monitor the behavior of the membrane dye (green) and the aqueous content (red) of the particles.
Upon lowering the pH to a value below the fusion pH, the particles will fuse with the membrane. Hemifusion, the merging of the outer leaflet of the viral membrane with the outer leaflet of the target membrane, will be visible as a sudden change in the green fluorescence of a particle. Upon the subsequent fusion of the two remaining distal leaflets a pore will be formed and the red-emitting fluorophore in the viral particle will be released under the target membrane. This event will give rise to a decrease of the red fluorescence of individual particles. Finally, the integrated fluorescence from a pH-sensitive fluorophore that is embedded in the target membrane reports on the exact time of the pH drop.
From the three fluorescence-time traces, all the important events (pH drop, lipid mixing upon hemifusion, content mixing upon pore formation) can now be extracted in a straightforward manner and for every particle individually. By collecting the elapsed times for the various transitions for many individual particles in histograms, we can determine the lifetimes of the corresponding intermediates. Even hidden intermediates that do not have a direct fluorescent observable can be visualized directly from these histograms.

We present an in vitro, two-color fluorescence assay to visualize the fusion of single virus particles with a fluid target bilayer. By labeling viral particles with fluorophores that differentially stain the viral membrane and its interior, we are able to monitor the kinetics of hemifusion and pore formation.

Membrane fusion is an essential step during entry of enveloped viruses into cells. Conventional fusion assays typically report on a large number of fusion ...

Analysis of mRNA Nuclear Export Kinetics in Mammalian Cells by Microinjection

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although the nuclear export of mRNA has been studied extensively in Xenopus oocytes1 and genetically tractable organisms such as yeast2 and the Drosophila derived S2 cell line3, few studies had been conducted in mammalian cells. Furthermore the kinetics of mRNA export in mammalian somatic cells could only be inferred indirectly4,5. In order to measure the nuclear export kinetics of mRNA in mammalian tissue culture cells, we have developed an assay that employs the power of microinjection coupled with fluorescent in situ hybridization (FISH). These assays have been used to demonstrate that in mammalian cells, the majority of mRNAs are exported in a splicing dependent manner6,7, or in manner that requires specific RNA sequences such as the signal sequence coding region (SSCR) 6. In this assay, cells are microinjected with either in vitro synthesized mRNA or plasmid DNA containing the gene of interest. The microinjected cells are incubated for various time points then fixed and the sub-cellular localization of RNA is assessed using FISH. In contrast to transfection, where transcription occurs several hours after the addition of nucleic acids, microinjection of DNA or mRNA allows for rapid expression and allows for the generation of precise kinetic data.

Here we describe an assay that employs the power of microinjection coupled with fluorescent in situ hybridization in order to accurately measure the nuclear export kinetics of mRNA in mammalian somatic cells.

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although ...

Photobleaching Assays (FRAP &amp; FLIP) to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with good spatial and temporal resolution. Here we describe how to perform FRAP and FLIP assays of chromatin proteins, including H1 and HP1, in mouse embryonic stem (ES) cells. In a FRAP experiment, cells are transfected, either transiently or stably, with a protein of interest fused with the green fluorescent protein (GFP) or derivatives thereof (YFP, CFP, Cherry, etc.). In the transfected, fluorescing cells, an intense focused laser beam bleaches a relatively small region of interest (ROI). The laser wavelength is selected according to the fluorescent protein used for fusion. The laser light irreversibly bleaches the fluorescent signal of molecules in the ROI and, immediately following bleaching, the recovery of the fluorescent signal in the bleached area - mediated by the replacement of the bleached molecules with the unbleached molecules - is monitored using time lapse imaging. The generated fluorescence recovery curves provide information on the protein's mobility. If the fluorescent molecules are immobile, no fluorescence recovery will be observed. In a complementary approach, Fluorescence Loss in Photobleaching (FLIP), the laser beam bleaches the same spot repeatedly and the signal intensity is measured elsewhere in the fluorescing cell. FLIP experiments therefore measure signal decay rather than fluorescence recovery and are useful to determine protein mobility as well as protein shuttling between cellular compartments. Transient binding is a common property of chromatin-associated proteins. Although the major fraction of each chromatin protein is bound to chromatin at any given moment at steady state, the binding is transient and most chromatin proteins have a high turnover on chromatin, with a residence time in the order of seconds. These properties are crucial for generating high plasticity in genome expression1. Photobleaching experiments are therefore particularly useful to determine chromatin plasticity using GFP-fusion versions of chromatin structural proteins, especially in ES cells, where the dynamic exchange of chromatin proteins (including heterochromatin protein 1 (HP1), linker histone H1 and core histones) is higher than in differentiated cells2,3.

Photobleaching Assays FRAP &amp; FLIP to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

We describe photobleaching methods including Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) to monitor chromatin protein dynamics in embryonic stem (ES) cells. Chromatin protein dynamics, which is considered to be one of the means to study chromatin plasticity, is enhanced in pluripotent cells.

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with ...

In vitro Transcription and Capping of Gaussia Luciferase mRNA Followed by HeLa Cell Transfection

In vitro transcription is the synthesis of RNA transcripts by RNA polymerase from a linear DNA template containing the corresponding promoter sequence (T7, T3, SP6) and the gene to be transcribed (Figure 1A). A typical transcription reaction consists of the template DNA, RNA polymerase, ribonucleotide triphosphates, RNase inhibitor and buffer containing Mg2+ ions.
Large amounts of high quality RNA are often required for a variety of applications. Use of in vitro transcription has been reported for RNA structure and function studies such as splicing1, RNAi experiments in mammalian cells2, antisense RNA amplification by the "Eberwine method"3, microarray analysis4 and for RNA vaccine studies5. The technique can also be used for producing radiolabeled and dye labeled probes6. Warren, et al. recently reported reprogramming of human cells by transfection with in vitro transcribed capped RNA7. The T7 High Yield RNA Synthesis Kit from New England Biolabs has been designed to synthesize up to 180 &mu;g RNA per 20 &mu;l reaction. RNA of length up to 10kb has been successfully transcribed using this kit. Linearized plasmid DNA, PCR products and synthetic DNA oligonucleotides can be used as templates for transcription as long as they have the T7 promoter sequence upstream of the gene to be transcribed.
Addition of a 5' end cap structure to the RNA is an important process in eukaryotes. It is essential for RNA stability8, efficient translation9, nuclear transport10 and splicing11. The process involves addition of a 7-methylguanosine cap at the 5' triphosphate end of the RNA. RNA capping can be carried out post-transcriptionally using capping enzymes or co-transcriptionally using cap analogs. In the enzymatic method, the mRNA is capped using the Vaccinia virus capping enzyme12,13. The enzyme adds on a 7-methylguanosine cap at the 5' end of the RNA using GTP and S-adenosyl methionine as donors (cap 0 structure). Both methods yield functionally active capped RNA suitable for transfection or other applications14 such as generating viral genomic RNA for reverse-genetic systems15 and crystallographic studies of cap binding proteins such as eIF4E16.
In the method described below, the T7 High Yield RNA Synthesis Kit from NEB is used to synthesize capped and uncapped RNA transcripts of Gaussia luciferase (GLuc) and Cypridina luciferase (CLuc). A portion of the uncapped GLuc RNA is capped using the Vaccinia Capping System (NEB). A linearized plasmid containing the GLuc or CLuc gene and T7 promoter is used as the template DNA. The transcribed RNA is transfected into HeLa cells and cell culture supernatants are assayed for luciferase activity. Capped CLuc RNA is used as the internal control to normalize GLuc expression.

In vitro Transcription and Capping of Gaussia Luciferase mRNA Followed by HeLa Cell Transfection

This method describes high yield in vitro synthesis of both capped and uncapped mRNA from a linearized plasmid containing the Gaussia luciferase (GLuc) gene. The RNA is purified and a fraction of the uncapped RNA is enzymatically capped using the Vaccinia virus capping enzyme. In the final step, the mRNA is transfected into HeLa cells and cell culture supernatants are assayed for luciferase activity.

In vitro transcription is the synthesis of RNA transcripts by RNA polymerase from a linear DNA template containing the corresponding promoter sequence ...

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 ...

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

Measuring the Osmotic Water Permeability Coefficient (Pf) of Spherical Cells: Isolated Plant Protoplasts as an Example

Studying AQP regulation mechanisms is crucial for the understanding of water relations at both the cellular and the whole plant levels. Presented here is a simple and very efficient method for the determination of the osmotic water permeability coefficient (Pf) in plant protoplasts, applicable in principle also to other spherical cells such as frog oocytes. The first step of the assay is the isolation of protoplasts from the plant tissue of interest by enzymatic digestion into a chamber with an appropriate isotonic solution. The second step consists of an osmotic challenge assay: protoplasts immobilized on the bottom of the chamber are submitted to a constant perfusion starting with an isotonic solution and followed by a hypotonic solution. The cell swelling is video recorded. In the third step, the images are processed offline to yield volume changes, and the time course of the volume changes is correlated with the time course of the change in osmolarity of the chamber perfusion medium, using a curve fitting procedure written in Matlab (the &#8216;PfFit&#8217;), to yield Pf.

Measuring the Osmotic Water Permeability Coefficient Pf of Spherical Cells: Isolated Plant Protoplasts as an Example

Measuring the osmotic water permeability coefficient (Pf) of cells can help understand the regulatory mechanisms of aquaporins (AQPs). Pf determination in spherical plant cell protoplasts presented here involves protoplasts isolation and numerical analysis of their initial rate of volume change as a result of an osmotic challenge during constant bath perfusion.

Studying AQP regulation mechanisms is crucial for the understanding of water relations at both the cellular and the whole plant levels. Presented here is ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...