Samir Hamdan aided in the development of this technique. E. coli proteins are from the lab of Prof. Nick Dixon, University of Wollongong, and T7 proteins are from Prof. Charles Richardson, Harvard Medical School. Work is supported by the National Institutes of Health (GM077248 to A.M.v.O.) and the Jane Coffin Childs Foundation (J.J.L.).

Before performing a single-molecule replication experiment, a few materials need to be prepared in advance. 
1. DNA Replication Template
The substrate for the replication reaction is a biotinylated, tailed M13 rolling circle prepared using standard molecular biology techniques1,2.
Materials: M13mp18 single-stranded DNA, Biotinylated tail oligonucleotide primer (5'-Biotin-T36AATTCGTAATCATGGTCATAGCTGTTTCCT-3'), T7 DNA polymerase, Heat block, Phenol/Isoamyl Alcohol/Chloroform
<ol>
 <li>Anneal the biotinylated tail oligo to the M13 by adding a 10-fold excess of oligo in TBS buffer (330 nM tail oligo, 33 nM M13).</li>
 <li>Heat the mixture to 65 &deg;C in the heat block. Once heated, turn the heat block off and allow slow cooling to properly anneal the primer.</li>
 <li>Add the primed M13 to a mixture of 64 nM T7 DNA polymerase and T7 replication buffer containing dNTPs and MgCl2.</li>
 <li>Incubate the reaction at 37 &deg;C for 12 minutes and quench with 100 mM EDTA.</li>
 <li>Purify the filled-in product DNA with phenol/chloroform/isoamyl alcohol extraction and dialyze into TE buffer or other suitable storage buffer and determine DNA concentration ( Back-extract each organic phase to recover any residual primed M13).</li>
 <li>One iteration of template preparation can easily make several milliliters of nanomolar concentration template, enough for hundreds to thousands of single-molecule experiments.</li>
</ol>
2. Functionalized Coverslips
For attaching the DNA to the glass coverslip, the glass is first functionalized with an aminosilane, which is then coupled to biotinylated PEG molecules. This coating helps to reduce the nonspecific interactions of DNA and replication proteins with the surface1,3.
Materials: Glass coverslips, Staining jars, 3-aminopropyltriethoxysilane, Methoxy-PEG5k-NHS ester, Biotin-PEG5k-NHSester, Acetone, 1M KOH, Ethanol, Oven, Bath sonicator
<ol>
 <li>Thoroughly clean the glass coverslips by placing them in staining jars and filling the jars with ethanol. Sonicate for 30 minutes and rinse out the jars and fill with 1 M KOH. Sonicate for 30 minutes Repeat both washes.</li>
 <li>For the first functionalization reaction, all traces of water need to be removed. Fill the jars with acetone and sonicate for 10 min. Rinse the jars again with acetone.</li>
 <li>Prepare a 2 % v/v solution of the silane reagent in acetone. Add to the staining jars and agitate for 2 minutes. This reaction couples the alkoxy group of the aminosilane to the glass, leaving a reactive amine for the next coupling step. Quench the reaction with a large excess of water. </li>
 <li>Dry the coverslips by baking them at 110 &deg;C in the oven for 30 minutes. </li>
 <li>Prepare a 50:1 methoxy:biotin PEG mixture in 100 mM NaHCO3 pH 8.2. Aim for around 0.2 % w/v biotinylated PEG. </li>
 <li>Pipette 100 &mu;L of the PEG mixture onto a dry silanized coverslip and place another coverslip on top. Including a glass coverslip spacer will allow separation of the coverslips.</li>
 <li>Incubate the coverslips in the PEG solution for 3 hours, then separate each pair of coverslips and wash extensively with water. Be careful to keep the coverslips functionalized side up as only once side will be coated with PEG.</li>
 <li>Dry the coverslips and store under vacuum. The surfaces remain stable for at least a month, so dozens of coverslips can be made in a batch and used as needed.</li>
</ol>
These materials should be made ahead of time, and then used for each experiment. To start every single-molecule experiment, begin by assembling the flow chamber.
3. Flow Chamber Preparation
The experiment is performed using a simple flow chamber constructed with a functionalized coverslip, double-sided tape, a quartz slide and tubing. One flow chamber is prepared for each single-molecule experiment1,4.
Materials: Double-sided tape, Razor blade, Quartz slide with holes for tubing, Quick-dry epoxy, Functionalized coverslip (see above), Streptavidin solution (25 &mu;L of 1 mg/mL in PBS), Tubing, Blocking buffer (20 mM Tris pH 7.5, 2 mM EDTA, 50 mM NaCl, 0.2 mg/mL BSA, 0.005% Tween-20)
<ol>
 <li>Begin by placing 20-25 mL of blocking buffer in a desiccator to remove any air bubbles for later steps.</li>
 <li>Take a PEG-functionalized coverslip and spread 125 of PBS-diluted streptavidin solution over the surface. Leave this to incubate for 20 minutes while preparing other parts of the chamber, allowing streptavidin to bind surface biotins.</li>
 <li>Cut a piece of double-sided tape to match the size of the quartz slide. Mark the position of the tubing holes on the tape with a pencil so that an outline of the flow chamber can be drawn on the tape (1 mg/mL).</li>
 <li>Make a 2 mm-wide rectangle around the holes. This will serve as the flow channel, so be sure to make straight sides and leave room around the tubing inlet and outlet. If desired, multiple channels or different sizes can be used.</li>
 <li>Cut along the drawn outline, making straight, clean cuts to make sure no adhesive protrudes into the channel. </li>
 <li>Clean the quartz thoroughly using acetone or ethanol to remove adhesive from the previous flow cell construction.</li>
 <li>Find the best alignment of the channel outline, remove one side of the adhesive backing and carefully place the tape onto the quartz slide. Be careful to align the tape properly, as the inlet and outlet holes need to remain unblocked.</li>
 <li>Cut lengths of tubing for the inlet and outlet of the flow cell. It helps to cut the end of the tube at about a 30&deg; angle so that the tube will not press flat against the chamber bottom. Place the tubes on some sort of support to suspend them for easy attachment to the flow cell in the next steps.</li>
 <li>Rinse the streptavidin-coated coverslip thoroughly with water and dry using compressed air. Remember that only one side is functionalized; be careful not to turn the coverslip over. Remove the other side of the tape backing and place the quartz slide onto the coverslip.</li>
 <li> Lightly press on the coverslip to push out any air trapped in the adhesive. This will help avoid any air bubbles getting into the flow channel.</li>
 <li> Seal the sides of the chamber with epoxy. Insert the cut tubing into the holes of the quartz and seal in place with epoxy. This needs to dry for a few minutes.</li>
 <li> Once dry, begin blocking the surface by pulling some of the degassed blocking buffer through one of tubes. A 21-gauge needle fits perfectly inside the 0.76 mm tubing. Flush out a few times to remove air bubbles, and allow the chamber to incubate for at least half an hour.</li>
</ol>
4. Single-Molecule Replication Experiment
Materials: Prepared flow chamber, SYTOX Orange, TIRF microscope, 532 nm laser, CCD camera, Computer with image acquisition software, Rolling-circle DNA substrate, Replication proteins
<ol>
<li>After the flow cell has been blocked, everything is ready to begin the single-molecule experiment.</li>
<li>Take the flow cell and place it on the microscope stage. Hold the chamber in place with stage clips and be sure that the flow channel is positioned straight along the y-axis.</li>
<li>Connect the flow cell outlet tube to the syringe pump using a larger diameter connector tubing or a needle. Place the inlet in blocking buffer and pull back on the syringe to remove any air in the tube. A gentle flick of the outlet tube will help clear any air bubbles trapped in the flow cell.</li>
<li>Dilute the stock DNA template to 25 pM in 1 mL of blocking buffer. Flow into the chamber at moderate flow rate to allow good surface coverage of DNA. 0.025 mL/min for 30 minutes works well. This can be varied based on how many DNA molecules are on the surface for each batch of coverslips or how many are desired for the experiment.</li>
<li>Once DNA is in, wash out excess DNA using blocking buffer. Wash for at least 200 flow-cell volumes to get rid of all the free DNA.</li>
<li>Begin degassing the replication buffer. For the E. coli replication experiments, do this at 37 &deg;C to reduce the risk of bubbles in the heated flow chamber.</li>
<li>Turn on the laser and camera. The cooled CCD needs to reach temperature before it can be used. </li>
<li>If performing a 37 &deg;C experiment, turn the heater on. Be sure the objective is in contact with the flow cell or it will be a heat sink later.</li>
<li>After washing and degassing, the reaction can be prepared. Mix nucleotides, DTT, and SYTOX in the replication buffer. Then add proteins and mix gently.</li>
<li>Flow the reaction mixture into the flow cell. This flow speed needs to be sufficient to stretch dsDNA. For a 2-mm wide, 100-&mu;m tall flow channel 0.04 mL/min works well.</li>
<li> Allow time for the mixture to enter the chamber and begin imaging. A good tip is to move the field of view to the side of the channel and focus on the adhesive material. This will ensure you are near the surface for focusing. </li>
<li> Adjust laser power, microscope focus, and TIRF angle. Keep the power low to avoid any photocleavage of the stained DNA. Acquire at 1-5 frames/second for several minutes depending on the rate of DNA replication. If desired, repeat to get longer replication trajectories or move to a new field to see more molecules.</li>
<li> After the reaction mixture is almost empty, it is a good idea to flow in more buffer with SYTOX to see other fields of view. Taking multiple images of different replicated molecules provides statistics for processivity determination.</li>
</ol>
5. Representative Results
Actively replicating molecules are easily seen as growing lines of stained DNA. In our experiments, flowing 25 pM of the M13 substrate gives 100-1000 molecules in a 60x, 125 &mu;m x 125 &mu;m field of view (Figure 1). Performing replication experiments using the E. coli replisome proteins at 37 &deg;C (see Materials for details) gives 5-50 replicating molecules per field of view. At equivalent concentrations of the T7 replisome, which initiates on the substrate much more efficiently, we observe &gt;70% of the molecules in a field replicate. These conditions yield a product density so high that individual molecules are difficult to resolve and analyze, so the T7 experiments are performed at lower protein concentrations.
Data Analysis: To obtain rate data, simply plot the endpoint of the DNA vs. time and calculate the slope (Figure 2). For processivity, determine the total length of the DNA. Both of these numbers will need to be converted to base pairs. Before performing the experiment, you should know the pixel size of the camera at the proper magnification. For basepair conversion, a good estimation is simply converting based on the crystallographic length of DNA, 2.9 bp/nm. However, laminar flow will not completely stretch the DNA, so a DNA length calibration should be performed by taking a known length DNA, e.g. 48,502 bp &lambda; DNA, attaching it to the flow cell with a biotinylated oligo and measuring its SYTOX-stained length at the flow rate used for replication experiment. Determining the number of base pairs/pixel will allow accurate calculation of replication rates and processivities. Taking numerous images and movies will provide large numbers of product molecule, allowing plotting of rate and processivity distributions (Figure 2)1.
<img src="/files/ftp_upload/1529/1529fig1.jpg" alt="Figure 1" /> Figure 1: (From Ref. 1) a) Cartoon of the rolling-circle replication assay. (SA, streptavidin). Leading-strand synthesis extends the tail linking the circle and surface. The tail is converted to dsDNA by the lagging strand polymerase and stretched by laminar flow. 
b) Example field of view with SYTOX Orange and 532 nm excitation. Small fluorescent spots are tethered, non-replicated substrates. Note the extreme length of the products and the large number of products per field (each flow cell contains &gt;5000 such fields).
<img src="/files/ftp_upload/1529/1529fig2.jpg" alt="Figure 2" /> Figure 2: (adapted from Ref. 1) a,b) Kymographs of typical replicating molecules from T7 (a) and E. coli (b) experiments. c) Endpoint trajectories of molecules from a) and b) plotted vs. time. Trajectories are fitted with linear regression to obtain replication rates. Shown examples are 99.4 bp s-1 and 467.1 bp s-1. d) Length distributions of replication products, fit with single exponential decay to obtain processivity: 25.3 &plusmn; 1.7 kbp (T7), 85.3 &plusmn; 6.1 kbp (E. coli). e) Rate distributions of single molecule trajectories, fit with single Gaussians. Means: 75.9 &plusmn; 4.8 bp s-1 (T7), 535.5 &plusmn; 39 bp s-1 (E. coli).

<ol>
	<li>Tanner, N. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+single-molecule+observation+of+rolling-circle+DNA+replication.">Real-time single-molecule observation of rolling-circle DNA replication.</a> Nucleic Acids Res. 37, e27 (2009).</li><li>Tabor, S., Huber, H. E., Richardson, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Escherichia+coli+thioredoxin+confers+processivity+on+the+DNA+polymerase+activity+of+the+gene+5+protein+of+bacteriophage+t7.">Escherichia coli thioredoxin confers processivity on the DNA polymerase activity of the gene 5 protein of bacteriophage t7.</a> J Biol Chem. 262, 16212-16223 (1987).</li><li>Sofia, S. J., Premnath, V. V., Merrill, E. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Poly(ethylene+oxide)+grafted+to+silicon+surfaces:+Grafting+density+and+protein+adsorption.">Poly(ethylene oxide) grafted to silicon surfaces: Grafting density and protein adsorption.</a> Macromolecules. 31, 5059-5070 (1998).</li><li>Tanner, N. A., van Oijen, A. M., Vengrova, S., Dalgaard, J. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+observation+of+prokaryotic+DNA+replication.">Single-molecule observation of prokaryotic DNA replication.</a> DNA replication: Methods and protocols. 521, 397-410 (2009).</li><li>Hamdan, S. M., Loparo, J. J., Takahashi, M., Richardson, C. C., van Oijen, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+DNA+replication+loops+reveal+temporal+control+of+lagging-strand+synthesis.">Dynamics of DNA replication loops reveal temporal control of lagging-strand synthesis.</a> Nature. 457, 336-339 (2009).</li><li>Heller, R. C., Marians, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Replication+fork+reactivation+downstream+of+a+blocked+nascent+leading+strand.">Replication fork reactivation downstream of a blocked nascent leading strand.</a> Nature. 439, 557-562 (2006).</li></ol>

One critical control is examining the effect of the stain, SYTOX Orange, on the replication proteins of interest. A simple way to do this is to perform the replication experiment in the flow cell as described but with the omission of SYTOX. After the reaction mixture has flowed through the chamber, add buffer with SYTOX to stain DNA and examine the length distribution of replicated molecules. Alternatively, standard bulk reactions can be used to check any effect of SYTOX on replication rate and efficiency.
The experiment described here uses only a single flow channel. This can be changed easily by creating multi-channel flow chambers or using PDMS or similar microfluidic devices. Increasing the number of channels greatly facilitates screening of protein concentrations, mutants, or inhibitor molecules and increases the rapidity of replication data collection.
As mentioned, we perform the E. coli replication experiments at 37 &deg;C using a home-built aluminum flow cell heater, a resistive heating element (cartridge heater) and variable power supply. This gives good temperature stability and avoids the purchase of an objective heater. To calibrate the heater, we simply drilled a hole in the center of a quartz flow cell top, inserted a thermocouple into the flow channel and flowed buffer as normal. Measuring the flow cell buffer temperature at increasing voltages allows accurate heating.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>M13mp18 ssDNA</td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>N4040</td>
<td>&nbsp;</td>
<tr>
<td>Biotinylated Tail Oligo</td>
<td>&nbsp;</td>
<td>Integrated DNA Technologies</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>T7 DNA Polymerase</td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>M0274</td>
<td>Use T7 replication buffer for substrate preparation</td>
</tr>
<tr>
<td>Phenol/ Isoamyl Alcohol/ Chloroform</td>
<td>&nbsp;</td>
<td>Roche</td>
<td>03117987001</td>
<td>24:24:1 v/v</td>
</tr>
<tr>
<td>3-aminopropyl-triethoxysilane</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>A3648</td>
<td>Other aminosilanes can be used or mixed with non-amine reactive silanes for sparser surfaces</td>
</tr>
<tr>
<td>Succinimidyl propionate PEG</td>
<td>&nbsp;</td>
<td>Nektar</td>
<td>&nbsp;</td>
<td>Similar PEGs can be purchased from Nanocs, CreativePEGWorks, etc.</td>
</tr>
<tr>
<td>Biotin-PEG-NHS</td>
<td>&nbsp;</td>
<td>Nektar</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Double-sided tape</td>
<td>&nbsp;</td>
<td>Grace BioLabs</td>
<td>SA-S-1L</td>
<td>100 &mu;m thickness</td>
</tr>
<tr>
<td>Quartz slide</td>
<td>&nbsp;</td>
<td>Technical Glass</td>
<td>20 mm (W)x 50 mm (L)x 1mm (H)</td>
<td>Size to fit on coverslips. Drill holes with diamond-tip drill bits (DiamondBurs.net)</td>
</tr>
<tr>
<td>Polyethylene tubing</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>427416</td>
<td>0.76 mm ID, 1.22 OD 
Other size tubing can be substituted.</td>
</tr>
<tr>
<td>Streptavidin</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>S4762</td>
<td>Make 1 mg/mL solution, 25 &mu;L aliquots in PBS pH 7.3</td>
</tr>
<tr>
<td>Deoxyribonucleotide triphosphate solution mix</td>
<td>&nbsp;</td>
<td>New England Biolabs</td>
<td>N0447</td>
<td>&nbsp;</td>
<tr>
<td>Ribonucleotide triphosphate solutions</td>
<td>&nbsp;</td>
<td>Amersham</td>
<td>272056 272066 272076 272086</td>
<td>&nbsp;</td>
<tr>
<td>SYTOX Orange</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>S11368</td>
<td>Other dsDNA stain can be used</td>
</tr>
<tr>
<td>Fluorescence Microscope with 60x TIRF objective</td>
<td>&nbsp;</td>
<td>Olympus</td>
<td>IX-71</td>
<td>Microscope, camera, etc. can be substituted for similar equipment</td>
</tr>
<tr>
<td>Syringe Pump</td>
<td>&nbsp;</td>
<td>Harvard Apparatus</td>
<td>11 Plus</td>
<td>Operate in refill mode to facilitate solution changes</td>
</tr>
<tr>
<td>532 nm laser</td>
<td>&nbsp;</td>
<td>Coherent</td>
<td>Compass 215M-75</td>
<td>Select wavelength to correspond to stain of choice</td>
</tr>
<tr>
<td>EMCCD Camera</td>
<td>&nbsp;</td>
<td>Hamamatsu</td>
<td>ImagEM</td>
<td>&nbsp;</td>
<tr>
<td>Emission filter</td>
<td>&nbsp;</td>
<td>Chroma</td>
<td>HQ600/75m</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>
			<div>
			T7 Replication: 40 mM Tris pH 7.5, 50 mM potassium glutamate, 10 mM magnesium chloride, 100 &mu;g/mL BSA, with 5 mM dithiothreitol, 600 &mu;M dNTPs, 300 &mu;M ATP, 300 &mu;M CTP, and 15 nM SYTOX Orange added immediately before use. Proteins added as: 5 nM gp4 (hexamer), 40 nM polymerase (1:1 gp5: thioredoxin), 360 nm gp2.51,5.
E. coli Replication: 50 mM HEPES pH 7.9, 12 mM magnesium acetate, 80 mM potassium chloride, 100 &mu;g/mL BSA with 10 mM dithiothreitol, 40 &mu;M dNTPs, 200 &mu;M rNTPs, and 15 nM SYTOX Orange added immediately before use. Proteins added as: 30 nM DnaB (hexamer), 180 nM DnaC (monomer), 30 nM &alpha;&epsilon;&theta;, 15 nM &tau;2&gamma;1&delta;&delta;&rsquo;&chi;&psi; or &tau;3&delta;&delta;&rsquo;&chi;&psi;, 30 nM &beta; (dimer), 300 nM DnaG, 250 nM SSB (tetramer), 20 nM PriA, 40 nM PriB, 320 nM PriC, 480 nM DnaT1,6		</div>

visualizing single-molecule dna replication with fluorescence microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


Watch this Scientific Journal Video about Visualizing Single-molecule DNA Replication with Fluorescence Microscopy at JoVE.com

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

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22.4K Views.  Harvard Medical School. This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

Video: Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, such as nucleic acids and proteins. Single molecule FRET (sm-FRET) measurements on immobilized molecules permit long observations of the system -effectively until both dyes photobleach- resulting in time-traces that report on biomolecular dynamics with a broad range of timescales from milliseconds to minutes. To facilitate the acquisition of large number of traces for statistical analyses, the process must be automated and the sample environment should be tightly controlled over the entire measurement time (~12 hours). This is accomplished using an automated scanning confocal microscope that allows the interrogation of thousands of single molecules overnight, and a microfluidic cell that permits the controlled exchange of buffer, with restricted oxygen content and maintains a constant temperature throughout the entire measuring period. Here we show how to assemble the microfluidic device and how to activate its surface for DNA immobilization. Then we explain how to prepare a buffer to maximize the photostability and lifetime of the fluorophores. Finally, we show the steps involved in preparing the setup for the automated acquisition of time-resolved single molecule FRET traces of DNA molecules.

In this article we describe how we obtain FRET traces from individual DNA molecules immobilized to a surface using an automated scanning confocal microscope.

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, ...

Direct Observation of Enzymes Replicating DNA Using a Single-molecule DNA Stretching Assay

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. Linearized &lambda; DNA is modified to have a biotin on the end of one strand, and a digoxigenin moiety on the other end of the same strand. The biotinylated end is attached to a functionalized glass coverslip and the digoxigeninated end to a small bead. The assembly of these DNA-bead tethers on the surface of a flow cell allows a laminar flow to be applied to exert a drag force on the bead. As a result, the DNA is stretched close to and parallel to the surface of the coverslip at a force that is determined by the flow rate (Figure 1). The length of the DNA is measured by monitoring the position of the bead. Length differences between single- and double-stranded DNA are utilized to obtain real-time information on the activity of the replication proteins at the fork. Measuring the position of the bead allows precise determination of the rates and processivities of DNA unwinding and polymerization (Figure 2).

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system.

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Cell Electrofusion Visualized with Fluorescence Microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

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 ...

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Few naturally-occurring plasmids are maintained in mammalian cells. Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus (EBV) and Kaposi's Sarcoma-associated herpesvirus (KSHV), which cause multiple human malignancies 1-3. These two genomes are replicated in a licensed manner, each using a single viral protein and cellular replication machinery, and are passed to daughter cells during cell division despite their lacking traditional centromeres 4-8.
Much work has been done to characterize the replications of these plasmid genomes using methods such as Southern blotting and fluorescence in situ hybridization (FISH). These methods are limited, though. Quantitative PCR and Southern blots provide information about the average number of plasmids per cell in a population of cells. FISH is a single-cell assay that reveals both the average number and the distribution of plasmids per cell in the population of cells but is static, allowing no information about the parent or progeny of the examined cell.
Here, we describe a method for visualizing plasmids in live cells. This method is based on the binding of a fluorescently tagged lactose repressor protein to multiple sites in the plasmid of interest 9. The DNA of interest is engineered to include approximately 250 tandem repeats of the lactose operator (LacO) sequence. LacO is specifically bound by the lactose repressor protein (LacI), which can be fused to a fluorescent protein. The fusion protein can either be expressed from the engineered plasmid or introduced by a retroviral vector. In this way, the DNA molecules are fluorescently tagged and therefore become visible via fluorescence microscopy. The fusion protein is blocked from binding the plasmid DNA by culturing cells in the presence of IPTG until the plasmids are ready to be viewed.
This system allows the plasmids to be monitored in living cells through several generations, revealing properties of their synthesis and partitioning to daughter cells. Ideal cells are adherent, easily transfected, and have large nuclei. This technique has been used to determine that 84% of EBV-derived plasmids are synthesized each generation and 88% of the newly synthesized plasmids partition faithfully to daughter cells in HeLa cells. Pairs of these EBV plasmids were seen to be tethered to or associated with sister chromatids after their synthesis in S-phase until they were seen to separate as the sister chromatids separated in Anaphase10. The method is currently being used to study replication of KSHV genomes in HeLa cells and SLK cells. HeLa cells are immortalized human epithelial cells, and SLK cells are immortalized human endothelial cells. Though SLK cells were originally derived from a KSHV lesion, neither the HeLa nor SLK cell line naturally harbors KSHV genomes11. In addition to studying viral replication, this visualization technique can be used to investigate the effects of the addition, removal, or mutation of various DNA sequence elements on synthesis, localization, and partitioning of other recombinant plasmid DNAs.

A method of observing individual DNA molecules in live cells is described. The technique is based on the binding of a fluorescently tagged lac repressor protein to binding sites engineered into the DNA of interest. This method can be adapted to follow many recombinant DNAs in live cells over time.

Few naturally-occurring plasmids are maintained in mammalian cells.  Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus  (EBV) ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Studying DNA Looping by Single-Molecule FRET

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into nucleosomes. Thermodynamics of dsDNA bending has been studied by a method called cyclization which relies on DNA ligase to covalently join short sticky ends of a dsDNA. However, ligation efficiency can be affected by many factors that are not related to dsDNA looping such as the DNA structure surrounding the joined sticky ends, and ligase can also affect the apparent looping rate through mechanisms such as nonspecific binding. Here, we show how to measure dsDNA looping kinetics without ligase by detecting transient DNA loop formation by FRET (Fluorescence Resonance Energy Transfer). dsDNA molecules are constructed using a simple PCR-based protocol with a FRET pair and a biotin linker. The looping probability density known as the J factor is extracted from the looping rate and the annealing rate between two disconnected sticky ends. By testing two dsDNAs with different intrinsic curvatures, we show that the J factor is sensitive to the intrinsic shape of the dsDNA.

This study presents a detailed experimental procedure to measure looping dynamics of double-stranded DNA using single-molecule Fluorescence Resonance Energy Transfer (FRET). The protocol also describes how to extract the looping probability density called the J factor.

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

Summary

Explore More Videos

Chapters in this video

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Direct Observation of Enzymes Replicating DNA Using a Single-molecule DNA Stretching Assay

Time-lapse Imaging of Mitosis After siRNA Transfection

Cell Electrofusion Visualized with Fluorescence Microscopy

Single-Molecule Imaging of Nuclear Transport

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

Studying DNA Looping by Single-Molecule FRET