The work was supported by NIH grants AI57159 (to S.C.H.) and AI72346 (to A.M.v.O.). S.C.H. is a Howard Hughes Medical Institute Investigator.

Glass cover slip functionalization
The planar bilayer used in the fusion assay is supported on a hydrated film of dextran. Dextran acts as a spacer between the planar bilayer and glass surface. This prevents membrane components from becoming stuck on the glass surface and also provides space in to which the contents of a virus particle can escape upon fusion. Glass coverslips are functionalized through treatment with an epoxy silane, which allows us to chemically bond dextran to the glass (Elender, et al. 1996).
<ol>
<li>Arrange 25 mm coverslips in a ceramic staining rack, and place the rack into a jar or beaker.</li>
<li>Fill the container with a 10% solution of laboratory grade glassware detergent. Place the container in an ultrasonic cleaner for 30 minutes.</li>
<li>Rinse the container and coverslip rack with deionized water, and refill with 1M postassium hydroxide. Return the container to the bath sonicator for another 30 minutes.</li>
<li>Repeat this cleaning procedure using acetone and ethanol.</li>
<li>Rinse the coverlips in water and dry.</li>
<li>Finally, clean the coverslips with an oxygen plasma stripper for three minutes.</li>
<li>The last cleaning step oxidizes the surface, making the glass uniformly hydrophilic and reactive in the following silanization steps.</li>
<li>Prepare a 0.2% solution of 3-glycidoxypropyltrimethoxy silane in isopropanol. Immerse the coverslips in this solution for five minutes.</li>
<li>Discard the silane solution and rinse the coverslips several times in isopropanol.</li>
<li>The coverslips are now coated with a layer of silane, but they must be cured to allow the silane to covalently bond to the glass.&nbsp; Place the coverslips in an oven set to 80 degrees Celsius for one hour.</li>
<li>While the coverslips are curing, weigh out dextran 500 and prepare a 30% w/v solution. Preheating the water will aid dissolution. We will use approximately 1 ml of this solution per coverslip. Once the dextran has dissolved, defoam the solution in a vacuum desiccator.</li>
<li>When the silanized coverslips have finished curing, remove them from the ceramic rack and arrange them flat on the inside of a plastic freezer box. Use a 1 ml pipette to cover the top surface of each coverslip with ~ 1 ml of the dextran solution. Close the freezer box and leave in a quiet place for 24-36 hours.</li>
<li>While grasping the coverslips with plastic forceps, pour off the excess dextran and replace them into the ceramic rack. Rinse the coverslips several times in water, and allow the coverslips to soak in water for 48 hours. The jar may be gently agitated on a table-top rotator.</li>
<li>Dry the coverslips in an oven set to 80 degrees Celsius and store in a vacuum desiccator.</li>
</ol>
Liposome Preparation
Supported lipid bilayers are formed by adsorbing liposomes to a dextran functionalized coverslip. The adsorbed liposomes fuse with eachother until membrane ruptures and spreads flat over the surface.
<ol>
<li>A typical liposome formulation used in the assay consists of 1,2,dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1-oleoyl-2-palmitoyl-sn-glycero-3-phosphocholine (POPC), cholesterol, bovine brain Disialoganglioside (GD1a), and N-((6-(biotinoyl)amino)hexanoyl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine in a ratio of 4:4:2:0.1:5x10-5. All components are stored in chloroform or chloroform and methanol solutions. </li>
<li>Prepare a mixture containing two micromoles of lipid by transferring volumes of each component into a glass test tube. Evaporate the solvent under a stream of argon or nitrogen gas. Remove the remaining solvent by leaving the test tube in a vacuum desiccator for two hours.</li>
<li>Resuspend the dried lipid film in 400 microliters of HNE buffer (5 mM HEPES, 145 mM NaCl, 0.1 mM EDTA). </li>
<li>Transfer the suspension to a plastic microcentrifuge tube. Freeze the suspension in a liquid nitrogen bath and thaw in a warm water bath. Repeat this freeze/thaw cycle four times.</li>
<li>Assemble a lipid extruder with a 100 nm polycarbonate membrane filter, and extrude the lipid suspension 25 times. The suspension should appear to be more transparent after extrusion.</li>
<li>Liposomes should be used on the day they are prepared. They may be stored at room temperature until used.</li>
</ol>
Microfluidic flow cell preparation
A simple microfluidic flow cell is made by sandwiching double stick tape between a quartz slide and a functionalized coverslip.
<ol>
<li>Obtain a 20x20x3 mm quartz slide. Use a diamond burr to drill two 1 mm holes on opposite sides.</li>
<li>Cut a 20 mm square from a sheet of double stick tape, and using a razor blade, cut out a 15 x 2 mm section from the center. </li>
<li>Peel off one side of the tape backing, and adhere the tape to the quartz slide. Adhere the other side to a functionalized coverslip.</li>
<li>Cut two 20 cm&nbsp; lengths of polyethylene tubing and insert them into the holes in the quartz slide.</li>
<li>Seal the flow cell with 5 minute epoxy glue.</li>
<li>When the glue has cured, prime the flow cell and tubing with buffer. </li>
<li>A supported lipid bilayer can be formed at this stage by drawing approximately 80 microliters of the liposome suspension into the channels.</li>
</ol>
Virus particle labeling
<ol>
<li>H3N2 influenza A (X-31) is typically used for the fusion assay, but other flu strains and other viruses have been used successfully.</li>
<li>Virus is labeled with up to two different fluorescent dyes to allow detection of hemifusion and pore-formation. </li>
<li>Dissolve sulforhodamine b (SRB) in HNE buffer to make a 20 mM solution. Combine 20 microliters of the dye solution with 10 microliters of the viral suspension, and leave at room temperature for 20 hours. During this period, dye will slowly accumulate inside the virus particles.</li>
<li>Remove the excess dye by passing the the virus suspension through a PD-10 desalting column. If enough viral material is used, a colored band can be seen in the column. This band contains labeled virus and can be collected as it is eluted.</li>
<li>If no band is visible, collect 200 microliter fractions and determine which fractions contain labeled virus by measuring 565 nm absorption on a spectrophotometer. The virus should elute in a total volume of approximately 0.8 ml. </li>
<li>Label the viral envelope by adding 13 microliters of a 2 mM dimethyl formamide (DMF) solution of octadecyl rhodamine 110 (Rh110C18) to the SRB labeled virus suspension.</li>
<li>Stir the suspension by attaching the tube to a laboratory rotator and leave for 3 hours.</li>
<li>Purify the virus from excess Rh110C18 using a PD-10 desalting column as described before.</li>
</ol>
Microscope configuration
The experiment is performed on a fluorescence microscope equipped with a high NA oil immersion objective suitable for total internal reflection fluorescence microscopy. The 488 and 568 nm lines from an argon/krypton gas laser are used to excite the Rh110C18 and SRB labeled virus. Simultaneous imaging of each label is accomplished by splitting the green and red fluorescence emission with a dichroic mirror and independently focusing the images onto separate halves of a electron multiplying CCD camera.
Performing the Assay
Execution of the fusion assay involves a stepwise assembly of the biochemical components within the flow cell: assembly of the planar lipid bilayer, docking of labeled virus particles, coating the surface with fluorescein, and initiation of the reaction with low pH buffer (Figure 1A).
<ol>
<li>Mount the flow-cell onto the microscope stage, and attach the outlet tubing to a syringe pump set to flow at a rate of 40 microliters per minute.</li>
<li>Clear the flow channels of excess liposomes by flowing in 300 microliters of HNE.</li>
<li>Focus the microscope and adjust the laser power to illuminate the surface with 350 W/cm2 of 488 nm light and 70 W/cm2 of 568 nm light. If you are using a 1.45 NA 60x oil immersion objective, set the laser intensities to 100 micro Watts and 20 micro Watts, respectively.</li>
<li>Pump labeled virus (diluted 1/100) into a flow cell. As the virus diffuses to the bottom surface of the flow channel, particles will begin to stick to the ganglioside incorporated in the&nbsp; lipid bilayer. </li>
<li>When a suitable density of docked virus particles has been attained (up to around 1000 particles per field of view). Switch the inlet tubing to a solution containing two micrograms per milliliter fluorescein labeled streptavidin. The labeled streptavidin will bind to biotin coupled lipid molecules in the bilayer, and eventually a dim green background will be visible.</li>
<li>Wash the flow channel with 300 microliters of HNE.</li>
<li>Choose an imaging area, focus the microscope and prepare the CCD camera for time-lapsed image acquisition. Set the exposure time to 100 ms and collect frames at a rate of 10 Hz.</li>
<li>Initiate fusion by flowing in an acidic buffer containing 10 mM sodium citrate and 140 mM NaCl. The pH of the buffer should be adjusted to be below the critical pH of the virus being assayed. X-31 fuses at pH&rsquo;s below 5.5.</li>
</ol>
Representative Results
Figure 1B shows snapshots of bilayer docked virions before and during fusion. Each diffraction limited spot represents an individual virus particle. Red and green fluorescence have been separately imaged onto the top and bottom halves of a CCD camera, allowing simultaneous observation of the viral envelope and content labels. There are typically twice as many spots in the green channel compared to the red, and this reflects the lower efficiency of labeling by the content dye compared to the envelope label. The green background fluorescence emitted from surface bound fluorescein disappears upon arrival of the citrate buffer and allows determination of the start time of the fusion reaction. Hemifusion events are detected as bursts of green fluorescence as the quenched Rh110C18 in the viral envelope diffuses across the hemifusion stalk and is diluted in the planar membrane. An outward expanding fluorescent cloud can be seen around hemifused particles, demonstrating free diffusion of the fatty-acyl linked dye in the planar membrane. Pore formation is observed as the decay of red fluorescence from SRB trapped within the interior of the viral particles. The opening of a fusion pore allows the dye to escape into the aqueous space below the planar bilayer and out of the observation region.
Fluorescence intensity trajectories for each particle, plotted from the integrated intensities of individual particles, facilitate determination of the yield and times of hemifusion and pore formation (Fig. 1C). Hemifusion and pore formation event times are determined as the point at which the maximum slope of a Rh110C18 fluorescence burst or SRB signal decay occurs. In a successful experiment, about 50 percent of the particles labeled with Rh110C18 yield dequenching signals, and 30 percent of the SRB labeled particles give useable signal. Of the particles labeled with both dyes, approximately 10 percent show both lipid mixing and pore formation signals.
Fig. 2A-C shows the distribution of hemifusion and pore formation lag times from a typical experiment. Fig.2D-F shows event distributions of events combined from five experiments performed under the same conditions. Even with a modest number of observations, the shape of the histograms is apparent. Because each experiment is synchronized by detection of the pH drop time, multiple experiments can be combined and detailed information about rate limiting intermediate steps can be obtained.
<img src="/files/ftp_upload/1484/1484fig1a.jpg" border="0" alt="figure 1" /> <img src="/files/ftp_upload/1484/1484fig1b.jpg" border="0" alt="figure 1" /> Figure 1. Experimental design. A) Virus particles are labeled with two fluorescent dyes to monitor the kinetics of hemifusion and fusion pore formation. Fluorescence is collected by a high-NA microscope objective and imaged onto a CCD. B) Fluorescence images before (left) and during (right) the fusion of individual viral particles. Top and bottom half of each image correspond to the red and green fluorescence, respectively, of the same ~ 50 x 100 mm2 area of the supported bilayer. C) The fluorescence intensity of the red SRB viral content tracer (upper trace), the green Rh110C18 membrane dye (middle trace), and the fluorescein pH sensor (lower trace) provide exact times elapsed between pH drop, hemifusion, and fusion. Please <a href="https://www.jove.com/files/ftp_upload/1484/1484fig1.jpg">click here</a> to see a larger version of figure 1.
<img border="0" src="/files/ftp_upload/1484/1484fig2tmb.jpg" alt="figure 2" /> Figure 2. Fusion kinetics of fluorescently labeled virus. Distributions of the time elapsed between the pH drop and hemifusion (A, D) and pore formation (B,E). The rise and decay of the distributions indicate the presence of multiple intermediate steps. The transition between hemifusion and opening of a fusion pore for individual particles is exponentially distributed, suggesting a single rate-limiting step. Panels A-C show the results of a single experiment. Panels D-F are compiled from five separate experiments performed under the same conditions. Please <a href="https://www.jove.com/files/ftp_upload/1484/1484fig2.jpg">click here</a> to see a larger version of figure 2.

<ol>
	<li>Elender, G., Kuhner, M., Sackmann, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functionalisation+of+Si/SiO2+and+glass+surfaces+with+ultrathin+dextran+films+and+deposition+of+lipid+bilayers.">Functionalisation of Si/SiO2 and glass surfaces with ultrathin dextran films and deposition of lipid bilayers.</a> Biosens Bioelectron. 11, 565-577 (1996).</li><li>Floyd, D. L., Ragains, J. R., Skehel, J. J., Harrison, S. 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=Single-particle+kinetics+of+influenza+virus+membrane+fusion.">Single-particle kinetics of influenza virus membrane fusion.</a> Proc Natl Acad Sci U S A. 105, 15382-15387 (2008).</li></ol>

Preparation of fluid and continuous supported lipid bilayers can be challenging. Trace amounts of contaminating material or surface defects will prevent spreading of bilayers. Careful cleaning and deposition of a uniform layer of dextran are essential.
Prolonged imaging of virus particles can bleach the fluorescent labels or cause them to become inactivated. Photo-damage of this kind is well known in the bio-imaging and single molecule fields and is generally believed to stem from the generat...

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		<div>
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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Corning cover glass squares, 25 mm</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>CLS286525</td>
<td>&nbsp;</td>
<tr>
<td>Fused silica slides</td>
<td>&nbsp;</td>
<td>Technical Glass</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Staining rack</td>
<td>&nbsp;</td>
<td>Thomas Scientific</td>
<td>8542E40</td>
<td>&nbsp;</td>
<tr>
<td>(3- glycidoxypropyl)trimethoxysilane</td>
<td>&nbsp;</td>
<td>Gelest</td>
<td>SIG5840.1</td>
<td>&nbsp;</td>
<tr>
<td>Dextran T-500</td>
<td>&nbsp;</td>
<td>Pharmacosmos</td>
<td>5510 0500 4006</td>
<td>&nbsp;</td>
<tr>
<td>1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC)</td>
<td>&nbsp;</td>
<td>Avanti</td>
<td>850375</td>
<td>&nbsp;</td>
<tr>
<td>1-palmitoyl-2oleoyl-sn-glycero-3-phosphocholine (POPC)</td>
<td>&nbsp;</td>
<td>Avanti</td>
<td>850457</td>
<td>&nbsp;</td>
<tr>
<td>Cholesterol</td>
<td>&nbsp;</td>
<td>Avanti</td>
<td>700000</td>
<td>&nbsp;</td>
<tr>
<td>N-((6-(biotinoyl)amino)hexanoyl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (biotin-X DHPE)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>B1616</td>
<td>&nbsp;</td>
<tr>
<td>Streptavidin, fluorescein conjugate</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>S-869</td>
<td>&nbsp;</td>
<tr>
<td>Disialoganglioside GD1a from bovine brain</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>G2392</td>
<td>&nbsp;</td>
<tr>
<td>Mini Extruder</td>
<td>&nbsp;</td>
<td>Avanti</td>
<td>610000</td>
<td>&nbsp;</td>
<tr>
<td>0.1 micron polycarbonate membrane filters</td>
<td>&nbsp;</td>
<td>Whatman</td>
<td>800309</td>
<td>&nbsp;</td>
<tr>
<td>10 mm drain discs (membrane supports)</td>
<td>&nbsp;</td>
<td>Whatman</td>
<td>230300</td>
<td>&nbsp;</td>
<tr>
<td>Sulforhodamine b sodium salt</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>S1402</td>
<td>&nbsp;</td>
<tr>
<td>Octadecyl rhodamine 110 (Rh110C18)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>See Floyd et al. 2008 for synthesis protocol</td>
</tr>
<tr>
<td>PD-10 desalting columns</td>
<td>&nbsp;</td>
<td>GE Healthcare</td>
<td>17-0851-01</td>
<td>&nbsp;</td>
<tr>
<td>Nikon fluorescence microscope</td>
<td>&nbsp;</td>
<td>Nikon</td>
<td>TE2000-U</td>
<td>&nbsp;</td>
<tr>
<td>Plan Apo TIRF 60x 1.45 NA objective</td>
<td>&nbsp;</td>
<td>Nikon</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Innova 70C Spectrum Ar/Kr laser</td>
<td>&nbsp;</td>
<td>Coherent</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Syringe Pump</td>
<td>&nbsp;</td>
<td>Harvard Apparatus</td>
<td>702213</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

method for measurement of viral fusion kinetics at the single particle level

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

Watch this Scientific Journal Video about Method for Measurement of Viral Fusion Kinetics at the Single Particle Level at JoVE.com

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

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

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

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12.8K Views.  Harvard Medical School. We present an in vitro, two-color fluorescence assay to visualize the fusion of single virus particles with a fluid target bilayer. By labeling viral particles with fluorophores that differentially stain the viral membrane and its interior, we are able to monitor the kinetics of hemifusion and pore formation.

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

Single Particle Electron Microscopy Reconstruction of the Exosome Complex Using the Random Conical Tilt Method

Single particle electron microscopy (EM) reconstruction has recently become a popular tool to get the three-dimensional (3D) structure of large macromolecular complexes. Compared to X-ray crystallography, it has some unique advantages. First, single particle EM reconstruction does not need to crystallize the protein sample, which is the bottleneck in X-ray crystallography, especially for large macromolecular complexes. Secondly, it does not need large amounts of protein samples. Compared with milligrams of proteins necessary for crystallization, single particle EM reconstruction only needs several micro-liters of protein solution at nano-molar concentrations, using the negative staining EM method. However, despite a few macromolecular assemblies with high symmetry, single particle EM is limited at relatively low resolution (lower than 1 nm resolution) for many specimens especially those without symmetry. This technique is also limited by the size of the molecules under study, i.e. 100 kDa for negatively stained specimens and 300 kDa for frozen-hydrated specimens in general.
For a new sample of unknown structure, we generally use a heavy metal solution to embed the molecules by negative staining. The specimen is then examined in a transmission electron microscope to take two-dimensional (2D) micrographs of the molecules. Ideally, the protein molecules have a homogeneous 3D structure but exhibit different orientations in the micrographs. These micrographs are digitized and processed in computers as "single particles". Using two-dimensional alignment and classification techniques, homogenous molecules in the same views are clustered into classes. Their averages enhance the signal of the molecule's 2D shapes. After we assign the particles with the proper relative orientation (Euler angles), we will be able to reconstruct the 2D particle images into a 3D virtual volume.
In single particle 3D reconstruction, an essential step is to correctly assign the proper orientation of each single particle. There are several methods to assign the view for each particle, including the angular reconstitution1 and random conical tilt (RCT) method2. In this protocol, we describe our practice in getting the 3D reconstruction of yeast exosome complex using negative staining EM and RCT. It should be noted that our protocol of electron microscopy and image processing follows the basic principle of RCT but is not the only way to perform the method. We first describe how to embed the protein sample into a layer of Uranyl-Formate with a thickness comparable to the protein size, using a holey carbon grid covered with a layer of continuous thin carbon film. Then the specimen is inserted into a transmission electron microscope to collect untilted (0-degree) and tilted (55-degree) pairs of micrographs that will be used later for processing and obtaining an initial 3D model of the yeast exosome. To this end, we perform RCT and then refine the initial 3D model by using the projection matching refinement method3.

This article describes a standard method to get a three-dimensional (3D) reconstruction of biological macromolecules using negative staining electron microscopy (EM). In this protocol, we explain how to get the 3D structure of the Saccharomyces cerevisiae exosome complex at medium resolution using the random conical tilt reconstruction method (RCT).

Single particle electron microscopy (EM) reconstruction has recently become a popular tool to get the three-dimensional (3D) structure of large ...

Measuring the Kinetics of mRNA Transcription in Single Living Cells

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

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

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

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

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

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

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

Analysis of Global RNA Synthesis at the Single Cell Level following Hypoxia

Hypoxia or lowering of the oxygen availability is involved in many physiological and pathological processes. At the molecular level, cells initiate a particular transcriptional program in order to mount an appropriate and coordinated cellular response. The cell possesses several oxygen sensor enzymes that require molecular oxygen as cofactor for their activity. These range from prolyl-hydroxylases to histone demethylases. The majority of studies analyzing cellular responses to hypoxia are based on cellular populations and average studies, and as such single cell analysis of hypoxic cells are seldom performed. Here we describe a method of analysis of global RNA synthesis at the single cell level in hypoxia by using Click-iT RNA imaging kits in an oxygen controlled workstation, followed by microscopy analysis and quantification.&nbsp; Using cancer cells exposed to hypoxia for different lengths of time, RNA is labeled and measured in each cell. This analysis allows the visualization of temporal and cell-to-cell changes in global RNA synthesis following hypoxic stress.

We describe a technique for analysis of global RNA synthesis in hypoxia using imaging. Click-chemistry labeling of RNA has not previously been performed under hypoxia and allows visualization of global RNA changes at the single cell level. This approach complements the existing averaged RNA techniques, allowing direct visualization of cell-to-cell changes in global RNA synthesis.

Hypoxia or lowering of the oxygen availability is involved in many physiological and pathological processes. At the molecular level, cells initiate a ...

Analysis of LINE-1 Retrotransposition at the Single Nucleus Level

Long interspersed nuclear element-1 (Line-1 or L1) accounts for approximately 17% of the DNA present in the human genome. While the majority of L1s are inactive due to 5' truncations, ~80-100 of these elements remain retrotransposition competent and propagate to different locations throughout the genome via RNA intermediates. While older L1s are believed to target AT rich regions of the genome, the chromosomal targets of newer, more active L1s remain poorly defined. Here we describe fluorescence in situ hybridization (FISH) methodology that can be used to track patterns of L1 insertion and rates of ectopic L1 incorporation at the single nucleus level. In these experiments, fluorescein isothiocyanate/cyanine-3 (FITC/CY3) labeled neomycin probes were employed to track L1 retrotransposition in vitro in HepG2 cells stably expressing ectopic L1. This methodology prevents errors in the estimation of rates of retrotransposition posed by toxicity and account for the occurrence of multiple insertions into a single nucleus.

Here we employ FISH methodology to track LINE-1 retrotransposition at the single nuclei level in chromosome spreads of HepG2 cell lines stably expressing synthetic LINE-1.

Long interspersed nuclear element-1 (Line-1 or L1) accounts for approximately 17% of the DNA present in the human genome. While the majority of L1s are ...

Open-source Single-particle Analysis for Super-resolution Microscopy with VirusMapper

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm allows for the routine imaging of nanoscale biological complexes and processes. This increase in resolution also means that methods popular in electron microscopy, such as single-particle analysis, can readily be applied to super-resolution fluorescence microscopy. By combining this analytical approach with super-resolution optical imaging, it becomes possible to take advantage of the molecule-specific labeling capacity of fluorescence microscopy to generate structural maps of molecular elements within a metastable structure. To this end, we have developed a novel algorithm &#8212; VirusMapper &#8212; packaged as an easy-to-use, high-performance, and high-throughput ImageJ plugin. This article presents an in-depth guide to this software, showcasing its ability to uncover novel structural features in biological molecular complexes. Here, we present how to assemble compatible data and provide a step-by-step protocol on how to use this algorithm to apply single-particle analysis to super-resolution images.

This manuscript uses the Fiji-based open-source software package VirusMapper to apply single-particle analysis to super-resolution microscopy images in order to generate precise models of nanoscale structure.

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm ...

Advanced Confocal Microscopy Techniques to Study Protein-protein Interactions and Kinetics at DNA Lesions

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological approach to analyzing protein kinetics at DNA lesions over time or protein-protein interactions on locally microirradiated chromatin. We also show how to recognize individual phases of the cell cycle using the Fucci cellular system to study cell-cycle-dependent protein kinetics at DNA lesions. A methodological description of the use of two UV lasers (355 nm and 405 nm) to induce different types of DNA damage is also presented. Only the cells microirradiated by the 405-nm diode laser proceeded through mitosis normally and were devoid of cyclobutane pyrimidine dimers (CPDs). We also show how microirradiated cells can be fixed at a given time point to perform immunodetection of the endogenous proteins of interest. For the DNA repair studies, we additionally describe the use of biophysical methods including FRAP (Fluorescence Recovery After Photobleaching) and FLIM (Fluorescence Lifetime Imaging Microscopy) in cells with spontaneously occurring DNA damage foci. We also show an application of FLIM-FRET (Fluorescence Resonance Energy Transfer) in experimental studies of protein-protein interactions.

Laser microirradiation is a useful tool for studies of DNA repair in living cells. A methodological approach for the use of UVA lasers to induce various DNA lesions is shown. We have optimized a method for local microirradiation that maintains the normal cell cycle; thus, irradiated cells proceed through mitosis.

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological ...

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, implantation of transgenic cells and tiny optical sensors. However, intravenous microinjections into small animals, which are mostly used in biological and veterinary laboratories, are very difficult and require trained personnel. Herein, we demonstrate a robust and efficient method for the introduction of microparticles into the circulatory system of adult zebrafish (Danio rerio) by injection into the fish kidney. To visualize the introduced microparticles in the vasculature, we propose a simple intravital imaging technique in fish gills. In vivo monitoring of the zebrafish blood pH was accomplished using an injected microencapsulated fluorescent probe, SNARF-1, to demonstrate one of the possible applications of the described technique. This article provides a detailed description of the encapsulation of pH-sensitive dye and demonstrates the principles of the quick injection and visualization of the obtained microcapsules for in vivo recording of the fluorescent signal. The proposed method of injection is characterized by a low mortality rate (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment. All described procedures can be performed on other small fish species, such as guppies and medaka.

This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, ...

Visualizing the Interaction Between the Qdot-labeled Protein and Site-specifically Modified &#955; DNA at the Single Molecule Level

The fluorescence microscopy&#160;has made great contributions in dissecting the mechanisms of complex biological processes at the single molecule level. In single molecule assays for studying DNA-protein interactions, there are two important factors for consideration: the DNA substrate with enough length for easy observation and labeling a protein with a suitable fluorescent probe. 48.5 kb &#955; DNA is a good candidate for the DNA substrate. Quantum dots (Qdots), as a class of fluorescent probes, allow long-time observation (minutes to hours) and high-quality image acquisition. In this paper, we present a protocol to study DNA-protein interactions at the single-molecule level, which includes preparing a site-specifically modified &#955; DNA and labeling a target protein with streptavidin-coated Qdots. For a proof of concept, we choose ORC (origin recognition complex) in budding yeast as a protein of interest and visualize its interaction with an ARS (autonomously replicating sequence) using TIRFM. Compared with other fluorescent probes, Qdots have obvious advantages in single molecule studies due to its high stability against photobleaching, but it should be noted that this property limits its application in quantitative assays.

Visualizing the Interaction Between the Qdot-labeled Protein and Site-specifically Modified &amp;#955; DNA at the Single Molecule Level

Here, we present a protocol to study DNA-protein interactions by total internal reflection fluorescence microscopy (TIRFM) using a site-specifically modified &#955; DNA substrate and a Quantum-dot labeled protein.

The fluorescence microscopy&#160;has made great contributions in dissecting the mechanisms of complex biological processes at the single molecule level. ...

Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

Centrioles are large macromolecular assemblies important for the proper execution of fundamental cell biological processes such as cell division, cell motility, or cell signaling. The green algae Chlamydomonas reinhardtii has proven to be an insightful model in the study of centriole architecture, function, and protein composition. Despite great advances toward understanding centriolar architecture, one of the current challenges is to determine the precise localization of centriolar components within structural regions of the centriole in order to better understand their role in centriole biogenesis. A major limitation lies in the resolution of fluorescence microscopy, which complicates the interpretation of protein localization in this organelle with dimensions close to the diffraction limit. To tackle this question, we are providing a method to purify and image a large number of C. reinhardtii centrioles with different orientations using super-resolution microscopy. This technique allows further processing of data through fluorescent single-particle averaging (Fluo-SPA) owing to the large number of centrioles acquired. Fluo-SPA generates averages of stained C. reinhardtii centrioles in different orientations, thus facilitating the localization of distinct proteins in centriolar sub-regions. Importantly, this method can be applied to image centrioles from other species or other large macromolecular assemblies.

Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

We have developed a strategy to purify and image a large number of centrioles in different orientations amenable for super-resolution microscopy and single-particle averaging.

Centrioles are large macromolecular assemblies important for the proper execution of fundamental cell biological processes such as cell division, cell ...