This work was supported by the Australian Research Council [DP11010246].

1. Blocking Replication with FROS
<ol>
	<li>Inducing the Replication Blockage
	<ol>
		<li>Dilute a fresh overnight culture of an E. coli strain carrying a tetO array and pKM118 to OD600nm = 0.01 in a dilute complex medium (0.1% tryptone, 0.05% yeast extract, 0.1% NaCl, 0.17 M KH2PO4, 0.72 M K2HPO4) with antibiotics as required for selection. 
		Note: Do not add tetracycline for selection as it is not compatible with this system. Make the volume of the culture equivalent to 10 ml per sample required. For example, the culture volume for the experimental design in Figure 1 is 60 ml.</li>
		<li>Grow at 30 &#176;C with shaking until OD600 nm = 0.05-0.1. Remove a 10 ml sample to serve as the uninduced control and add 0.1% arabinose to the remaining culture to induce the production of TetR-YFP from pKM1. Continue to grow both the uninduced and induced cultures. After 1 hr, check for the presence of a single focal point within each cell of the induced culture using fluorescence microscopy (see section 2).</li>
		<li>If the induced cells are confirmed blocked (more than 70% of cells have a single focus), record the OD600nm and take a 7.5 ml sample for analysis by 2-D gels (see section 3). Take an equivalent sample from the uninduced control culture.</li>
	</ol>
	</li>
	<li>Releasing the Replication Blockage
	<ol>
		<li>To release the replication block, add 100 ng ml-1 of the gratuitous inducer anhydrotetracycline (AT) to a 10 ml sample of the blocked cells. Continue to grow at 30 &#176;C for 10 min and take samples for cell density (1 ml), microscopy analysis (10 &#181;l) and neutral-neutral 2-dimensional agarose gels (7.5 ml).</li>
	</ol>
	</li>
	<li>Inducing Collapse of the Replisome
	<ol>
		<li>If the cells contain a temperature sensitive allele such as dnaBts or dnaCts that requires deactivation, move the flask containing the blocked cells to 42 &#176;C. Take samples for analysis at time points required (e.g., 1 hr after incubation). After the cells have incubated at 42 &#176;C for 1 hr, shift the cells to 30 &#176;C for 10 min (see Figure 1). 
		Note: The cells can be shifted to 30 &#176;C for restart analysis.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/54434/54434fig1.jpg" /> 
Figure 1: Overview of the FROS Replication Block and Release Experimental Procedure. E. coli strains carrying the tetO array are grown at 30 &#176;C. When the cells reach an OD600nm of above 0.05, TetR-YFP production is induced with arabinose (ara; 0.1%). Continue to grow a subpopulation of uninduced cells to act as controls at 30 &#176;C. A sample of the induced cells is analyzed via fluorescence microscopy after 1 - 2 hr (see 2.1). If replication is confirmed to be blocked, samples are taken for 2-D gel analysis indicated by the test tubes (see 3.1) and for viability tests (optional). The replication blockage can be removed with anhydrotetracycline (AT; 0.1 &#181;g/ml) and cells analyzed 10 min later. If cells carry a temperature sensitive allele (e.g. dnaBts), this can be inactivated by shifting the blocked cells to 42 &#176;C for appropriate analysis. <a href="https://www.jove.com/files/ftp_upload/54434/54434fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Fluorescence Microscopy
<ol>
	<li>Prepare an agarose pad by pipetting 500 &#181;l of molten agarose (1% in water) onto a microscope slide; overlay the coverslip immediately before the agarose solidifies. Store the slide(s) in wet tissues at 4 &#176;C until needed.</li>
	<li>When the agarose has solidified, remove the coverslip from the agarose pad and pipette 10 &#181;l of bacterial culture onto the center of the pad to prevent movement of the cells during visualization. Once the sample has dried (approximately 5 min), replace the coverslip. Place a drop of immersion oil onto the cover slip and place the slide under optics.</li>
	<li>Visualize the Cells Using Phase Microscopy at 100X Magnification.
	<ol>
		<li>On the Acquire dialog box within the imaging software, set exposure time to 100 msec. Capture the image by selecting Acquire. Do not move the stage. In the Acquire dialog box, select YFP as the illumination for the External Shutter Linked to Camera. Set the exporsure time to 1,000 msec. Turn off the stage light. Capture the fluorescence image by selecting Acquire. 
		Note: Time may vary depending upon light source, microscope and camera used.</li>
		<li>To overlay the phase and fluorescent images, select Color Combine from within the Display menu. In the Color Combine dialog box, select the YFP image as the green component and the phase image as the blue component. Click on color combine. Determine whether the population has had replication successfully blocked by establishing whether the majority of the cells have one focus per cell. 
		Note: Comparison to a sample from the control without added arabinose is useful to visually identify the difference in eYFP production.</li>
	</ol>
	</li>
</ol>
3. DNA Extraction/Agarose Plugs
<ol>
	<li>Add sodium azide (final concentration of 0.1%) to the 7.5 ml culture samples from steps 1.1.3 and 1.2.1. Incubate on ice for at least 5 min. 
	Note: Sodium azide is extremely toxic. Consult the Safety Data Sheet prior to commencing work and do not handle it until all safety precautions have been read and understood.</li>
	<li>Centrifuge the cells 5,000 x g for 10 min to pellet cells. Discard the supernatant and resuspend the pellet in 200 &#181;l of Pett IV (PIV) buffer (10 mM Tris:HCl (pH 8.0), 1 M NaCl). Microcentrifuge (13, 000 x g for 2 min) to pellet cells. Discard the supernatant. At this stage, the process the pellets further or store at -20 &#176;C.</li>
	<li>Resuspend the cells with the lowest cell density in 50 &#181;l PIV and place at 50 &#176;C. For all other samples, adjust the volumes of PIV so the final cell density is the same in each tube (e.g. Sample 1 (OD600nm = 0.128) resuspended in 50 &#181;l; Sample 2 (OD600nm = 0.138) resuspended in 54 &#181;l).
	<ol>
		<li>Add an equal volume of freshly prepared 0.8% agarose solution in PIV (final conc. 0.4%; e.g. Sample 1 50 &#181;l, Sample 2 54 &#181;l). Ensure that the samples, agarose and PIV are above 50 &#176;C to prevent solidification.</li>
	</ol>
	</li>
	<li>Treat a microscope slide with 40 &#181;l of an appropriate glass water-repellent or silicone solution. Rub the slide with a tissue until it is dry. 
	Note: This may be done in advance and the treated slides stored long term at room temperature.</li>
	<li>Place 20 &#181;l drops of the cell/agarose suspension on the slide to produce plugs that are hemispherical. 
	Note: The first sample (resuspended in 50 &#181;l PIV) will produce 5 plugs on one slide.</li>
	<li>When the plugs have solidified, slide them gently into a microfuge tube and add 1 ml cell lysis buffer (10 mM Tris-HCl [pH 8], 1 M NaCl, 100 mM Ethylenediaminetetraacetic acid (EDTA), 0.2% sodium deoxycholate, 0.5% N-Lauroylsarcosine sodium salt (Sarkosyl), 100 &#181;g ml&#8722;1 lysozyme, 50 &#181;g ml&#8722;1 RNase A). Incubate at 37 &#176;C for 2 hr.</li>
	<li>Remove the cell lysis buffer and add 1 ml EDTA/Sarcosyl/Proteinase K (ESP) solution (0.5 M EDTA, 1% N-Lauroylsarcosine sodium salt (Sarkosyl), 1 mg ml&#8722;1 proteinase K). Incubate at 50 &#176;C overnight or until the plugs are transparent. If a second overnight incubation is required, replace the ESP solution with fresh buffer after approximately 18 hr.</li>
	<li>Remove the ESP buffer and wash the plugs 5 times with 12 ml Tris-EDTA (TE) buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0), allowing a 30 min equilibration with each wash. Store at 4 &#176;C in 1 ml TE buffer.</li>
</ol>
4. Neutral-neutral 2-Dimensional Gel Electrophoresis
<ol>
	<li>Transfer an agarose plug from step 3.8 to a fresh microfuge tube and add 150 &#181;l of restriction enzyme buffer (1x concentration). Add 25-100 units of restriction enzyme (e.g., 60 units of EcoRV; (see Figure 3A)). Digest at 37 &#176;C for 6-8 hr.</li>
	<li>At 4 &#176;C, pour a 0.4% agarose gel (300 ml) in 1x Tris/Borate/EDTA (TBE) into a gel tray of approximately 25 x 25 cm. Insert a comb to make wells approximately the same width as the diameter of the plug. When the gel is set, remove the comb and move the gel to room temperature.</li>
	<li>Slide one of the digested agarose plugs into a well, positioning the flat slide of the plug against the side of the well where the DNA will enter the gel. Insert a single plug in the same manner into every second well.</li>
	<li>Pipette molten 0.4% agarose into the wells to seal the plugs in position.</li>
	<li>Load 1 kb DNA ladder in an empty well, again leaving a gap between the ladder and samples, and run the gel overnight (16 hr, 55 V) in 1x TBE at room temperature.</li>
	<li>Stain the gel in a water bath containing ethidium bromide (0.3 &#181;g ml-1) for 20 min. 
	Note: Ethidium bromide is considered a potent mutagen and a toxin. Consult the Safety Data Sheet prior to commencing work and do not handle it until all safety precautions have been read and understood.</li>
	<li>Visualize the DNA with a long-wave UV transilluminator and cut out each lane fragment with a straight, even cut, minimizing the inclusion of excess agarose on either side of the lane. 
	Note: For example, if the fragment of interest is 5.5 kb, cut a 7.5 cm fragment to include the DNA fragments from 5 kb to approximately 12 kb (see Figure 3A).</li>
	<li>Place the excised gel lane in a gel tray at 90&#176; to direction of DNA migration. 
	Note: Two rows of 3 gel fragments can fit in a 25 x 252 cm gel tray. The first row of gel lanes is at the top of the tray, the second row at 12.5 cm.</li>
	<li>Prepare 300 ml agarose for the second dimension gel (0.9% in 1x TBE with 0.3 &#181;g ml-1 ethidium bromide). When the agarose is cooled to 50 &#176;C, pipette the molten agarose around the gel slices to solidify their position. Pour the remaining agarose into the tray to a depth that is at least level with the gel slices.</li>
	<li>Once the gel is solidified, electrophorese at 4 &#176;C in 1x TBE containing 0.3 &#181;g ml-1 ethidium bromide at 220 V until the DNA has migrated approximately 10 cm. 
	Note: 4 hr is sufficient for a 5.5 kb fragment but a smaller fragment will need less time.</li>
	<li>Excise the blocks where the genomic DNA is present using the edge of a metal ruler, including the gel above it to include the DNA that is not visible (see Figure 3C).</li>
</ol>
5. Southern Hybridization
<ol>
	<li>Solution Preparation
	<ol>
		<li>Prepare Depurination buffer (0.125 M HCl solution). 
		Note: This buffer can be reused and is stable at room temperature for up to 1 month.</li>
		<li>Prepare Denaturation buffer (1.5 M NaCl, 0.5 M NaOH). 
		Note: Denaturation buffer can be reused once and is stable for up to 3 months at room temperature.</li>
		<li>Prepare Neutralization buffer (1.5 M NaCl, 0.5 M Tris). Adjust to pH 7.5 with HCl. 
		Note: Neutralization buffer can be reused once and is stable for up to 3 months at room temperature.</li>
		<li>Prepare 10x Saline Sodium Citrate (SSC) buffer (0.15 M Tri-sodium citrate, 1.5 M NaCl, pH is 7 - 8) for use in steps 5.2.4 and 5.2.6. From this buffer, make a 2x SSC stock (for use in step 5.2.9). 
		Note: 10x SSC and 2x SSC are stable at RT for 3 months.</li>
		<li>Prepare Modified Church and Gilbert Hybridization Buffer19 (0.5 M phosphate buffer [pH 7.2], 7% (w/v) sodium dodecyl sulfate (SDS), 10 mM EDTA). Leave at 65 &#176;C to dissolve thoroughly prior to use.</li>
	</ol>
	</li>
	<li>Transfer
	<ol>
		<li>Rinse the excised gel blocks in depurination solution for 10 min at RT on a rocker or orbital shaker. Keep the agitation speed low to avoid damaging the agarose blocks.</li>
		<li>Rinse the gel blocks briefly with distilled water and then with denaturation buffer for 30 min. Rinse again briefly in distilled water and then incubate the gel blocks in neutralization buffer for 30 min with gentle agitation at room temperature.</li>
		<li>Cut a nylon membrane to the exact size of the gel(s). Cut a nick in the top right hand corner to assist in orienting the membrane in future steps. 
		Note: Two gel blocks (6 samples) can be visualized on one membrane.</li>
		<li>Pour enough 10x SSC into a tray so that it will not dry out overnight. Create a platform approximately 5 cm above the level of the buffer. Saturate 3 sheets of 3MM chromatography paper with 10x SSC and place on the platform. Cut the chromatography paper so that it is long enough to be immersed in buffer at both ends and wide enough to fit the membrane gel on.</li>
		<li>Place the gel(s) on top of the saturated chromatography paper. Frame the gel with plastic wrap to prevent the buffer by-passing the gel during transfer. Carefully lay the cut membrane on top of the gel(s). Remove air bubbles between the membrane and gel by rolling a bottle or serological pipette gently across the membrane.</li>
		<li>Cut three sheets of 3MM chromatography paper to the same size as the membrane. Saturate the chromatography paper sheets with 10x SSC and place on top of the membrane. Remove any air bubbles that have formed.</li>
		<li>Stack paper towels at least 5 cm high on top of the blotting paper followed by a solid support (approximate weight of 1 kg for a 23 x 16 cm membrane). Allow the transfer to proceed overnight.</li>
		<li>Expose the membrane (DNA side up) to 1,200 J/m2 of UV to crosslink the DNA to the membrane. Do not rinse prior to this step.</li>
		<li>Rinse the membrane thoroughly in 2x SSC to prevent high background on the blot images. Use the blot immediately for hybridization (see 5.3) or dry it at RT, wrap in plastic wrap and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Hybridization
	<ol>
		<li>Preheat hybridization oven and bottles to 55 &#176;C.</li>
		<li>Add 100 &#181;g ml-1 bovine serum albumin (BSA) and 10 &#181;g ml-1 non-specific DNA (e.g. salmon sperm DNA) to the pre-warmed hybridization buffer.</li>
		<li>To allow for even coverage of the hybridization buffer when the membrane is rolled up, place the blot onto a similarly sized square of nylon mesh with the DNA-bound side of the membrane facing upwards. Roll the blot along its long edge. Slide the blot/mesh inside a hybridization bottle. Add an appropriate amount of pre-heated hybridization buffer to unroll the blot (e.g. 30 ml for a 23 x 16 cm2 membrane).</li>
		<li>Incubate in a pre-heated oven, rotating the bottles, for 1 hr.</li>
		<li>Prepare the probe by mixing 50 ng of purified PCR product homologous to the region of interest, 150 ng of random hexamer primers, buffer for Klenow fragment and H2O to make a volume of 13 &#181;l. Boil for 3 min then place immediately onto ice. Add 1 &#181;l of 1 mM dCTP/dGTP/dTTP mix, 1 &#181;l of Klenow fragment (5U) and 50 &#181;Ci deoxyadenosine 5&#39;-triphosphate radiolabeled on the alpha phosphate-(&#945;32P-dATP).</li>
		<li>Incubate at room temp for 1 hr. Add 1 &#181;l of 1 mM dNTP mix and 1 &#181;l of Klenow fragment. Incubate 20 min at RT.</li>
		<li>Boil probe for 5 min and add immediately to hybridization tubes. Hybridize overnight at 55 &#176;C, with rotation.</li>
		<li>Decant the probe from membrane. 
		Note: The probe is able to be reused once.</li>
		<li>Rinse the membrane briefly in pre-warmed, low stringency wash buffer (2x SSC, 0.1% (w/v) SDS). Add fresh low stringency wash buffer and incubate for 5 min at 55 &#176;C with rotation. Repeat.</li>
		<li>Wash twice in medium stringency wash buffer (1x SSC, 0.1% (w/v) SDS) for 10 min at 55 &#176;C with rotation.</li>
		<li>Wash twice in high stringency wash buffer (0.1x SSC, 0.1% (w/v) SDS) for 5 min at 55 &#176;C with rotation.</li>
		<li>Remove the membrane and dab with tissue to remove excess liquid. Wrap in plastic wrap ensuring there is no remaining moisture on the plastic wrap and place in exposure cassette with a pre-blanked storage phosphor screen. Close the cassette and expose for at least 2 hr before visualization using a phosphor imager.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

During chromosome duplication, the replication machinery will encounter various impediments that prevent its progress. To ensure the entire single-origin chromosome is replicated, bacteria have numerous pathways for repair of the DNA that then enables the replisome to be reloaded20,21. Lesions, single stranded breaks, double stranded breaks and proteins tightly bound to the DNA may each be dealt with using a dedicated pathway, although there is likely to be significant overlap in these pathways. The most commo...

During DNA replication, the replisome faces obstacles on the DNA that impair its progression. DNA damage including lesions and gaps as well as aberrant structures can prevent the replisome from proceeding1. Recently, it has been found that proteins bound to the DNA are the most common source of impediment to replication fork progression2. The knowledge of the events following the encounter of the replisome with a nucleoprotein block has previously been limited by the inability to induce such a block in the chromosome of a living cell at a known location. In vitro analysis has enhanced our understanding of the kinetic behavior of an active replisome when it meets a nucleoprotein blockage3, as well as the mechanistic details of the replisome itself4,5. Current understanding of the repair of replication is generally undertaken with UV as the damaging agent and studied using plasmid DNA in vivo6-8. While the proteins that may be involved in repair of DNA after it encounters an in vivo nucleoprotein block are generally understood from these studies, whether there are variations in the molecular events within the repair pathways owing to the distinct cause in the replication block is still yet to be determined.
Here, we describe a system that allows a nucleoprotein block to be established in a specific location of the chromosome using a Fluorescent Repressor Operator System (FROS). We utilize a strain of E. coli that has had an array of 240 tetO sites incorporated into the chromosome9. Each tetO site within the array has a 10 bp random sequence flanking it to increase the stability of the array by preventing RecA-mediated recombination within the array. This array, and variations of it, were originally used to understand E. coli chromosome dynamics10,11 but were then adapted to prevent in vivo replication12. The array has been found to be stably maintained and to block close to 100% of replication forks when bound by TetR10,12. The use of the similar lacO array in vitro has found as few as 22 sites were sufficient to block 90% of replication, although this shorter array was less effective in vivo13. To adapt the array to create a nucleoprotein blockage, the repressor protein must be highly overproduced under optimized conditions where it then binds to the array to create a roadblock. The formation of the blockage, and its subsequent release, can be monitored through the use of fluorescence microscopy if a fluorescently tagged variant of the Tet repressor is used. The status of replication in each cell is indicated by the number of foci seen, where a single focal point means only one copy of the array is present within the cell and multiple foci are indicative of active replication. This active replication is enabled when the nucleoprotein blockage is reversed by the addition of the gratuitous inducer that decreases the binding affinity of TetR for the operator site sufficiently for the replisome to proceed through the array. The repressor protein is still able to bind to the DNA with high enough affinity that the now multiple copies of the array can be visualized.
More intricate details of the events at the nucleoprotein blockage can be discovered using neutral-neutral two-dimensional agarose gel electrophoresis and Southern hybridization14-16. These techniques allow the analysis of DNA structures across the population. The replication intermediates that are formed during the event, and potentially remain unrepaired, can be visualized. By varying the restriction enzyme and probe utilized, the intermediates can be visualized not only in the array region but also upstream of the array when the replication fork regresses17,18. The regression takes place subsequent to the replisome dissociation; the leading and lagging nascent strands separate from the template strands and anneal to each other as the template strands concurrently re-anneal resulting in a four-way DNA structure (a Holliday junction).
Using this system it has been shown that the replication fork is not stable when it encounters this block18. In addition, temperature sensitive derivatives of replisome components can be utilized to prevent reloading of the replication fork once it has collapsed. Once a block is established, the strain can be shifted to a non-permissive temperature to ensure a synchronous deactivation of the replisome and a controlled prevention of reloading. This temperature-induced deactivation ensures all of the forks within the population have collapsed at a given time and allows the assessment of what happens when the replisome collapses, how the DNA is processed, and what is required to restart the process of DNA replication.
An advantage of the system described here is that the nucleoprotein block is fully reversible; therefore, the ability of the cells to recover from the nucleoprotein block is able to be followed. The addition of anhydrotetracycline to the cells will relieve the tight binding of the repressor, allowing a replication fork to proceed through and the cell to regain viability. The relief of the blockage can be visualized by neutral-neutral two-dimensional agarose gel electrophoresis after 5 min, and by microscopy within 10 min. Furthermore, viability analysis can reveal the ability of the strain to recover from the replication blockage and continue to proliferate.
By altering the genetic background of the strains used in the experimental procedure described here, the repair pathways for this type of blockage can be elucidated.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Tryptone</td>
			<td>Sigma-Aldrich</td>
			<td>16922</td>
			<td>Growth media component</td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>VWR</td>
			<td>27810.364</td>
			<td>Growth media component</td>
		</tr>
		<tr>
			<td>Yeast extract</td>
			<td>Sigma-Aldrich</td>
			<td>92144</td>
			<td>Growth media component</td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic</td>
			<td>Sigma-Aldrich</td>
			<td>P9791</td>
			<td>Growth media component; Potassium buffer component</td>
		</tr>
		<tr>
			<td>Potassium phosphate dibasic</td>
			<td>Sigma-Aldrich</td>
			<td>P3786</td>
			<td>Growth media component; Potassium buffer component</td>
		</tr>
		<tr>
			<td>L-Arabinose</td>
			<td>Sigma-Aldrich</td>
			<td>A3256</td>
			<td>For induction of TetR-YFP production</td>
		</tr>
		<tr>
			<td>Anhydrotetracycline hydrochloride</td>
			<td>Sigma-Aldrich</td>
			<td>37919</td>
			<td>Release of replication bloackage</td>
		</tr>
		<tr>
			<td>Axioskop 2 Fluorescence microscope</td>
			<td>Zeiss</td>
			<td>452310</td>
			<td>Visualization of cells</td>
		</tr>
		<tr>
			<td>eYFP filter set</td>
			<td>Chroma Technology</td>
			<td>41028</td>
			<td>Visualization of YFP</td>
		</tr>
		<tr>
			<td>CCD camera</td>
			<td>Hamamatsu</td>
			<td>Orca-AG</td>
			<td>Visualization of cells</td>
		</tr>
		<tr>
			<td>MetaMorph&#160; Software (Molecular Devices)</td>
			<td>SDR Scientific</td>
			<td>31282</td>
			<td>Version 7.8.0.0 used in the preparation of this manuscript</td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Bioline</td>
			<td>BIO-41025</td>
			<td>For agarose plugs and gel electrophoresis</td>
		</tr>
		<tr>
			<td>Original Glass Water Repellent (Rain-X)</td>
			<td>Autobarn</td>
			<td>DIO1470</td>
			<td>For agarose plug manufacture</td>
		</tr>
		<tr>
			<td>TRIS</td>
			<td>VWR</td>
			<td>VWRC103157P</td>
			<td>TE, TBE buffer component</td>
		</tr>
		<tr>
			<td>Ethylene diaminetetraacetic acid</td>
			<td>Ajax Finechem</td>
			<td>AJA180</td>
			<td>0.5 M EDTA disodium salt solution adjusted to pH 8.0 with NaOH.</td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Sigma-Aldrich</td>
			<td>S2002</td>
			<td>Bacteriostatic agent</td>
		</tr>
		<tr>
			<td>Hydrochloric acid</td>
			<td>Sigma-Aldrich</td>
			<td>258148</td>
			<td>TE buffer component</td>
		</tr>
		<tr>
			<td>Sodium deoxycholate</td>
			<td>Sigma-Aldrich</td>
			<td>D6750</td>
			<td>Cell lysis buffer component</td>
		</tr>
		<tr>
			<td>N-Lauroylsarcosine sodium salt (Sarkosyl)</td>
			<td>Sigma-Aldrich</td>
			<td>L5125</td>
			<td>Cell lysis and ESP buffer component</td>
		</tr>
		<tr>
			<td>Rnase A</td>
			<td>Sigma-Aldrich</td>
			<td>R6513</td>
			<td>Cell lysis buffer component</td>
		</tr>
		<tr>
			<td>Lysozyme</td>
			<td>Amresco</td>
			<td>6300</td>
			<td>Cell lysis buffer component</td>
		</tr>
		<tr>
			<td>Proteinase K</td>
			<td>Amresco</td>
			<td>AM0706</td>
			<td>ESP buffer component</td>
		</tr>
		<tr>
			<td>Sub-Cell Model 192 Cell</td>
			<td>BioRad</td>
			<td>1704507</td>
			<td>Electrophoresis system</td>
		</tr>
		<tr>
			<td>UV transilluminator 2000</td>
			<td>BioRad</td>
			<td>1708110</td>
			<td>Visualization of DNA</td>
		</tr>
		<tr>
			<td>Ethidium Bromide</td>
			<td>BioRad</td>
			<td>1610433</td>
			<td>Visualization of DNA</td>
		</tr>
		<tr>
			<td>Boric acid</td>
			<td>VWR</td>
			<td>PROL20185.360</td>
			<td>TBE component</td>
		</tr>
		<tr>
			<td>Hybond-XL nylon memrbane</td>
			<td>Amersham</td>
			<td>RPN203S</td>
			<td>Zeta-Probe Memrbane (BioRad 1620159) can also be used</td>
		</tr>
		<tr>
			<td>3MM Whatman chromatography paper</td>
			<td>GE Healthcare Life Sciences</td>
			<td>3030690</td>
			<td>Southern blotting</td>
		</tr>
		<tr>
			<td>HL-2000 Hybrilinker</td>
			<td>UVP</td>
			<td>95-0031-01/02</td>
			<td>Crosslinking of DNA and hybridization</td>
		</tr>
		<tr>
			<td>Deoxyribonucleic acid from salmon sperm</td>
			<td>Sigma-Aldrich</td>
			<td>31149</td>
			<td>Hybridization buffer component</td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Sigma-Aldrich</td>
			<td>S5881</td>
			<td>Denaturation buffer component</td>
		</tr>
		<tr>
			<td>Trisodium citrate dihydrate</td>
			<td>VWR</td>
			<td>PROL27833.363</td>
			<td>Transfer buffer</td>
		</tr>
		<tr>
			<td>Sodium dodecyl sulphate (SDS)</td>
			<td>Amresco</td>
			<td>227</td>
			<td>Wash buffer component</td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A7906</td>
			<td>Hybridization buffer component</td>
		</tr>
		<tr>
			<td>Random Hexamer Primers</td>
			<td>Bioline</td>
			<td>BIO-38028</td>
			<td></td>
		</tr>
		<tr>
			<td>Klenow fragment</td>
			<td>New England BioLabs</td>
			<td>M0212L</td>
			<td></td>
		</tr>
		<tr>
			<td>dNTP Set</td>
			<td>Bioline</td>
			<td>BIO-39025</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine 5&#8217;-triphosphate-32P-ATP</td>
			<td>PerkinElmer</td>
			<td>BLU502A</td>
			<td></td>
		</tr>
		<tr>
			<td>Storage Phosphor Screen</td>
			<td>GE Healthcare Life Sciences</td>
			<td>GEHE28-9564-76</td>
			<td>BAS-IP MS 3543 E multipurpose standard 35x43cm screen</td>
		</tr>
		<tr>
			<td>Typhoon FLA 7000</td>
			<td>GE Healthcare Life Sciences</td>
			<td>28-9558-09</td>
			<td>Visualization of blot</td>
		</tr>
		<tr>
			<td>Hybridization bottle</td>
			<td>UVP</td>
			<td>07-0194-02</td>
			<td>35 x 300mm</td>
		</tr>
	</tbody>
</table>

inducing a site specific replication blockage in e. coli using a fluorescent repressor operator system

Obstacles present on DNA, including tightly-bound proteins and various lesions, can severely inhibit the progression of the cell&#39;s replication machinery. The stalling of a replisome can lead to its dissociation from the chromosome, either in part or its entirety, leading to the collapse of the replication fork. The recovery from this collapse is a necessity for the cell to accurately complete chromosomal duplication and subsequently divide. Therefore, when the collapse occurs, the cell has evolved diverse mechanisms that take place to restore the DNA fork and allow replication to be completed with high fidelity. Previously, these replication repair pathways in bacteria have been studied using UV damage, which has the disadvantage of not being localized to a known site. This manuscript describes a system utilizing a Fluorescence Repressor Operator System (FROS) to create a site-specific protein block that can induce the stalling and collapse of replication forks in Escherichia coli. Protocols detail how the status of replication can be visualized in single living cells using fluorescence microscopy and DNA replication intermediates can be analyzed by 2-dimensional agarose gel electrophoresis. Temperature sensitive mutants of replisome components (e.g. DnaBts) can be incorporated into the system to induce a synchronous collapse of the replication forks. Furthermore, the roles of the recombination proteins and helicases that are involved in these processes can be studied using genetic knockouts within this system.

The FROS is an inducible, site-specific nucleoprotein block that enables replication intermediates to be visualized in living cells12,18. A general experimental design for sampling cells is illustrated in Figure 1. The timing of the sampling and variations in genetic background make this a versatile system for studying the repair of such a block. The schematic illustrates how temperature sensitive mutants, such as dnaBts and dnaCts that have b...

Watch this Scientific Journal Video about Inducing a Site Specific Replication Blockage in E. coli Using a Fluorescent Repressor Operator System at JoVE.com

Inducing a Site Specific Replication Blockage in E. coli Using a Fluorescent Repressor Operator System

We describe here a system utilizing a site-specific, reversible in vivo protein block to stall and collapse replication forks in Escherichia coli. The establishment of the replication block is evaluated by fluorescence microscopy and neutral-neutral 2-dimensional agarose gel electrophoresis is used to visualize replication intermediates.

Inducing a Site Specific Replication Blockage in E. coli Using a Fluorescent Repressor Operator System

Obstacles present on DNA, including tightly-bound proteins and various lesions, can severely inhibit the progression of the cell&#39;s replication ...

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8.9K Views.  University of Newcastle. We describe here a system utilizing a site-specific, reversible in vivo protein block to stall and collapse replication forks in Escherichia coli. The establishment of the replication block is evaluated by fluorescence microscopy and neutral-neutral 2-dimensional agarose gel electrophoresis is used to visualize replication intermediates.

Video: Inducing a Site Specific Replication Blockage in E. coli Using a Fluorescent Repressor Operator System

Transformation of Plasmid DNA into E. coli Using the Heat Shock Method

Transformation of plasmid DNA into E. coli using the heat shock method is a basic technique of molecular biology. It consists of inserting a foreign plasmid or ligation product into bacteria. This video protocol describes the traditional method of transformation using commercially available chemically competent bacteria from Genlantis. After a short incubation in ice, a mixture of chemically competent bacteria and DNA is placed at 42&deg;C for 45 seconds (heat shock) and then placed back in ice. SOC media is added and the transformed cells are incubated at 37&deg;C for 30 min with agitation. To be assured of isolating colonies irrespective of transformation efficiency, two quantities of transformed bacteria are plated. This traditional protocol can be used successfully to transform most commercially available competent bacteria. The turbocells from Genlantis can also be used in a novel 3-minute transformation protocol, described in the instruction manual.

Transformation of plasmid DNA into E. coli using the heat shock method is a basic technique of molecular biology. It consists of inserting a foreign ...

Principles of Site-Specific Recombinase (SSR) Technology

Site-specific recombinase (SSR) technology allows the manipulation of gene structure to explore gene function and has become an integral tool of molecular biology. Site-specific recombinases are proteins that bind to distinct DNA target sequences. The Cre/lox system was first described in bacteriophages during the 1980's. Cre recombinase is a Type I topoisomerase that catalyzes site-specific recombination of DNA between two loxP (locus of X-over P1) sites. The Cre/lox system does not require any cofactors. LoxP sequences contain distinct binding sites for Cre recombinases that surround a directional core sequence where recombination and rearrangement takes place. When cells contain loxP sites and express the Cre recombinase, a recombination event occurs. Double-stranded DNA is cut at both loxP sites by the Cre recombinase, rearranged, and ligated ("scissors and glue"). Products of the recombination event depend on the relative orientation of the asymmetric sequences.
SSR technology is frequently used as a tool to explore gene function. Here the gene of interest is flanked with Cre target sites loxP ("floxed"). Animals are then crossed with animals expressing the Cre recombinase under the control of a tissue-specific promoter. In tissues that express the Cre recombinase it binds to target sequences and excises the floxed gene. Controlled gene deletion allows the investigation of gene function in specific tissues and at distinct time points. Analysis of gene function employing SSR technology --- conditional mutagenesis -- has significant advantages over traditional knock-outs where gene deletion is frequently lethal.

Principles of Site-Specific Recombinase SSR Technology

The advent of site-specific recombinase (SSR) technology and the Cre/lox system has led to numerous advances in molecular biology, and has proven itself as a valuable tool for assessing gene function in transgenic animals. This interview discusses the mechanism of site specific recombination by Cyclization recombinase (Cre) and how the use of this enzyme has led to the development of conditional mutagenesis, which has significant advantages over traditional knock out strategies.

Site-specific recombinase (SSR) technology allows the manipulation of gene structure to explore gene function and has become an integral tool of molecular ...

Visualization of UV-induced Replication Intermediates in E. coli using Two-dimensional Agarose-gel Analysis

Inaccurate replication in the presence of DNA damage is responsible for the majority of cellular rearrangements and mutagenesis observed in all cell types and is widely believed to be directly associated with the development of cancer in humans. DNA damage, such as that induced by UV irradiation, severely impairs the ability of replication to duplicate the genomic template accurately. A number of gene products have been identified that are required when replication encounters DNA lesions in the template. However, a remaining challenge has been to determine how these proteins process lesions during replication in vivo. Using Escherichia coli as a model system, we describe a procedure in which two-dimensional agarose-gel analysis can be used to identify the structural intermediates that arise on replicating plasmids in vivo following UV-induced DNA damage. This procedure has been used to demonstrate that replication forks blocked by UV-induced damage undergo a transient reversal that is stabilized by RecA and several gene products associated with the RecF pathway. The technique demonstrates that these replication intermediates are maintained until a time that correlates with the removal of the lesions by nucleotide excision repair and replication resumes.

Visualization of UV-induced Replication Intermediates in E. coli using Two-dimensional Agarose-gel Analysis

We present a procedure by which two-dimensional agarose-gel analysis can be used to identify the structure of replication intermediates that occur following UV irradiation.

Inaccurate replication in the presence of DNA damage is responsible for the majority of cellular rearrangements and mutagenesis observed in all cell types ...

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular signatures, or to evolve proteins in the laboratory. Here, we present a protocol that allows the generation of large (>1011) mutant libraries for a given target sequence. This method is based on replication of a ColE1 plasmid encoding the desired sequence by a low-fidelity variant of DNA polymerase I (LF-Pol I). The target plasmid is transformed into a mutator strain of E. coli and plated on solid media, yielding between 0.2 and 1 mutations/kb, depending on the location of the target gene. Higher mutation frequencies are achieved by iterating this process of mutagenesis. Compared to alternative methods of mutagenesis, our protocol stands out for its simplicity, as no cloning or PCR are involved. Thus, our method is ideal for mutational labeling of plasmids or other Pol I templates or to explore large sections of sequence space for the evolution of activities not present in the original target. The tight spatial control that PCR or randomized oligonucleotide-based methods offer can also be achieved through subsequent cloning of specific sections of the library. Here we provide protocols showing how to create a random mutant library and how to establish drug-based selections in E. coli to identify mutants exhibiting new biochemical activities.

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Here we demonstrate a simple protocol to create a random mutant library for a given target sequence. We show how this method, which is performed in vivo in Escherichia coli, can be coupled with functional selections to evolve new enzymatic activities.

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular ...

A 3D System for Culturing Human Articular Chondrocytes in Synovial Fluid

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not regenerate very efficiently in vivo; and as a result, osteoarthritis leads to irreversible cartilage loss and is accompanied by chronic pain and immobility 1,2. Cartilage tissue engineering offers promising potential to regenerate and restore tissue function. This technology typically involves seeding chondrocytes into natural or synthetic scaffolds and culturing the resulting 3D construct in a balanced medium over a period of time with a goal of engineering a biochemically and biomechanically mature tissue that can be transplanted into a defect site in vivo 3-6. Achieving an optimal condition for chondrocyte growth and matrix deposition is essential for the success of cartilage tissue engineering. 
 In the native joint cavity, cartilage at the articular surface of the bone is bathed in synovial fluid. This clear and viscous fluid provides nutrients to the avascular articular cartilage and contains growth factors, cytokines and enzymes that are important for chondrocyte metabolism 7,8. Furthermore, synovial fluid facilitates low-friction movement between cartilaginous surfaces mainly through secreting two key components, hyaluronan and lubricin 9 10. In contrast, tissue engineered cartilage is most often cultured in artificial media. While these media are likely able to provide more defined conditions for studying chondrocyte metabolism, synovial fluid most accurately reflects the natural environment of which articular chondrocytes reside in.
 Indeed, synovial fluid has the advantage of being easy to obtain and store, and can often be regularly replenished by the body. Several groups have supplemented the culture medium with synovial fluid in growing human, bovine, rabbit and dog chondrocytes, but mostly used only low levels of synovial fluid (below 20&#37;) 11-25. While chicken, horse and human chondrocytes have been cultured in the medium with higher percentage of synovial fluid, these culture systems were two-dimensional 26-28. Here we present our method of culturing human articular chondrocytes in a 3D system with a high percentage of synovial fluid (up to 100&#37;) over a period of 21 days. In doing so, we overcame a major hurdle presented by the high viscosity of the synovial fluid. This system provides the possibility of studying human chondrocytes in synovial fluid in a 3D setting, which can be further combined with two other important factors (oxygen tension and mechanical loading) 29,30 that constitute the natural environment for cartilage to mimic the natural milieu for cartilage growth. Furthermore, This system may also be used for assaying synovial fluid activity on chondrocytes and provide a platform for developing cartilage regeneration technologies and therapeutic options for arthritis.

A 3D system of culturing human articular chondrocytes in high levels of synovial fluid is described. Synovial fluid reflects the most natural microenvironment for articular cartilage, and can be easily obtained and stored. This system thus can be used for studying cartilage regeneration and for screening therapeutics for treating arthritis.

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not ...

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

The Logic, Experimental Steps, and Potential of Heterologous Natural Product Biosynthesis Featuring the Complex Antibiotic Erythromycin A Produced Through E. coli

The heterologous production of complex natural products is an approach designed to address current limitations and future possibilities. It is particularly useful for those compounds which possess therapeutic value but cannot be sufficiently produced or would benefit from an improved form of production. The experimental procedures involved can be subdivided into three components: 1) genetic transfer; 2) heterologous reconstitution; and 3) product analysis. Each experimental component is under continual optimization to meet the challenges and anticipate the opportunities associated with this emerging approach.
Heterologous biosynthesis begins with the identification of a genetic sequence responsible for a valuable natural product. Transferring this sequence to a heterologous host is complicated by the biosynthetic pathway complexity responsible for product formation. The antibiotic erythromycin A is a good example. Twenty genes (totaling &gt;50 kb) are required for eventual biosynthesis. In addition, three of these genes encode megasynthases, multi-domain enzymes each ~300 kDa in size. This genetic material must be designed and transferred to E. coli for reconstituted biosynthesis. The use of PCR isolation, operon construction, multi-cystronic plasmids, and electro-transformation will be described in transferring the erythromycin A genetic cluster to E. coli.
Once transferred, the E. coli cell must support eventual biosynthesis. This process is also challenging given the substantial differences between E. coli and most original hosts responsible for complex natural product formation. The cell must provide necessary substrates to support biosynthesis and coordinately express the transferred genetic cluster to produce active enzymes. In the case of erythromycin A, the E. coli cell had to be engineered to provide the two precursors (propionyl-CoA and (2S)-methylmalonyl-CoA) required for biosynthesis. In addition, gene sequence modifications, plasmid copy number, chaperonin co-expression, post-translational enzymatic modification, and process temperature were also required to allow final erythromycin A formation.
Finally, successful production must be assessed. For the erythromycin A case, we will present two methods. The first is liquid chromatography-mass spectrometry (LC-MS) to confirm and quantify production. The bioactivity of erythromycin A will also be confirmed through use of a bioassay in which the antibiotic activity is tested against Bacillus subtilis. The assessment assays establish erythromycin A biosynthesis from E. coli and set the stage for future engineering efforts to improve or diversify production and for the production of new complex natural compounds using this approach.

The Logic, Experimental Steps, and Potential of Heterologous Natural Product Biosynthesis Featuring the Complex Antibiotic Erythromycin A Produced Through E. coli

The heterologous biosynthesis of erythromycin A through E. coli includes the following experimental steps: 1) genetic transfer; 2) heterologous reconstitution; and 3) product analysis. Each step will be explained in the context of the motivation, potential, and challenges in producing therapeutic natural products using E. coli as a surrogate host.

The heterologous production of complex natural products is an approach designed to address current limitations and future possibilities.  It is ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Detection of Intracellular Gene Expression in Live Cells of Murine, Human and Porcine Origin Using Fluorescence-labeled Nanoparticles

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, rat, human, pig and others. The verification of iPS clones mainly relies on the detection of the endogenous expression of different pluripotency genes. These genes mostly represent transcription factors which are located in the cell nucleus. Traditionally, the proof of their endogenous expression is supplied by immunohistochemical staining after fixation of the cells. This approach requires replicate cultures of each clone at this early stage to preserve validated clones for further experiments. The present protocol describes an approach with gene-specific nanoparticles which allows the evaluation of intracellular gene expression directly in live cells by fluorescence. The nanoparticles consist of a central gold particle coupled to a capture strand carrying a sequence complementary to the target mRNA as well as a quenched reporter strand. These nanoparticles are actively endocytosed and the target mRNA displaces the reporter strand which then start to fluoresce. Therefore, specific target gene expression can be detected directly under the microscope. In addition, the emitted fluorescence allows the identification, isolation and enrichment of cells expressing a specific gene by flow cytometry. This method can be applied directly to live cells in culture without any manipulation of the target cells.

This manuscript describes a novel technique which allows the detection of intracellular gene expression after endocytosis of fluorescence-labeled nanoparticles directly in live cells. The method does not require manipulation of the cells and is not restricted with respect to the target gene or species.

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, ...