Name: automated system for single molecule fluorescence measurements of surface-immobilized biomolecules
Uploaded: 2009-11-02T16:05:00
Duration: 10 min 57 s
Description: Watch this Scientific Journal Video about Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules at JoVE.com

We thank Chandran Sabanayagam for his help in developing the system, Joshua Edel for advice on designing the microfluidic device and the staff at the Rowland Institute at Harvard University for machining parts of the setup. We acknowledge useful discussions with members of Meller&#x2019;s group at the Rowland Institute and at Boston University, and members of T. Ha&#x2019;s group at UIUC for helpful information. Finally, we appreciate the financial support from the National Science Foundation (PHY-0701207) and the National Institute of Health (GM075893).

1 - Assembling the microfluidic cell
The microfluidic cell is made by fusing a patterned, 2 mm thick, polydimethylsiloxane (PDMS) disk to a freshly cleaned quartz cover slip. The PDMS disk contains four 30 &mu;l rectangular chambers that are accessed by inlet and outlet flexible capillaries connected to a buffer reservoir and a syringe pump, respectively, for the controlled flow of buffer. The &#xFB02;ow cell is supported on its back side by a glass window and placed on a custom-made cell holder containing a water cooled thermoelectric element to regulate temperature.
<ol>
 <li>To make the patterned PDMS disk, mix 10 parts silicone elastomer base with 1 part curing agent, mix well and leave degassing for 1 hour in vacuum.</li>
 <li>Gently pour the elastomer mix onto the microfluidic cell mold. In our case, this is a 1" silicon wafer with four strips (1x104 x 1x103 x 300 &mu;m) made from negative photoresist. Leave this to cure on a hot plate at 70&deg; C for two hours.</li>
 <li>Carefully peel the cured PDMS disk from the mold using a razor blade. Fuse the disk onto a 1'' glass window (containing 8 x 2 mm-diameter pre-drilled holes at both ends of the channels) by plasma oxidizing both surfaces for 30 seconds. Fuse the glass windows onto the flat side of the PDMS disk (the side not containing the channels).</li>
 <li>Insert a 2-inch long flexible fused silica capillary tubing through each glass window's hole until it reaches the channel. Inserting a needle first and then following with the capillary may facilitate this process. Seal the glass window holes using a fast curing silicone casting compound such as Kwik-Cast.</li>
 <li>Rinse the channels with ethanol and water, and plasma-activate the cell together with a freshly cleaned, 1-inch circular quartz cover slip for 30 seconds. We suggest cleaning this cover slip using piranha*. Fuse the cover slip to the side of the cell containing the channels, sealing the chambers.</li>
 <li>Leave the assembled cell to cure overnight at room temperature.</li>
</ol>
* Piranha is a solution of H2O2 and H2SO4 in 1:2.5 proportions. To clean the cover slips, immerse them in piranha at 90&deg; C for 20 min.
2 - Activating the surface for molecule immobilization
For nucleic acid studies, the simplest surface immobilization scheme consists of first coating a glass or quartz cover slip with biotinylated bovine serum albumin (BSA) and then with streptavidin. This allows the biotinylated sample to be immobilized with high specificity for days at room temperature. 
<ol>
 <li>Connect flexible tubing to the capillaries for easy buffer injection using a syringe and needle.</li>
 <li>Inject a solution of 0.1 mg/ml of biotinylated BSA in buffer A (10 mM TRIS-HCl, pH 8.0, 50 mM NaCl, filtered using 0.02 &mu;m filters) into the channel and incubate for at least 30 min.</li>
 <li>Flow 300-500 &mu;l of buffer A to remove any free biotinylated BSA from the channel, being careful not to trap any air bubbles inside the chamber.</li>
 <li>Inject a solution of 0.1 mg/ml of streptavidin in buffer A and incubate for 15 min. Then repeat Step 3.</li>
 <li>Flow through a 3-5 pM solution of the biotinylated sample. Perform a surface scan to check the coverage of fluorescent molecules. If necessary, increase the sample concentration to obtain better coverage. Incubate for 10 min and repeat Step 3.</li>
</ol>
The microfluidic cell is mounted on a motorized x-y stage allowing coarse (up to 25 mm) motion using optically encoded DC motors, and &#xFB01;ne (1 nm) motion using two closed-loop piezo actuators (Physik Instrumente model M-014 or similar). This can be done either before or after activating the surface for sample immobilization.
3 - Preparing the Imaging Buffer (IB)
The IB consists of an enzymatic oxygen scavenging system and a triplet quencher, such as Trolox. Since the IB is flushed through constantly overnight, at least 10 ml should be prepared in advance.
<ol>
 <li>Prepare 10 ml of 0.8% w/v D-glucose, at least 35 mM TRIS-HCl pH 8.0 and 50 mM NaCl in deionized water. The higher concentration of buffer is important for two reasons: dissolving Trolox will acidify the solution, and the enzymatic reaction will also lower the pH of the solution over time. Since this solution has a long shelf life it can be stored as a stock solution.</li>
 <li>Dissolve 5 mg of Trolox in the buffer prepared in Step 1 by vortexing for a couple of minutes and then shaking for &gt;10 min. Trolox is not very soluble in water at pH &gt; 7, so do not rush this step. Filter the solution using a 0.2 &mu;m filter and leave degassing in vacuum for 10 minutes.</li>
 <li>Prepare the "gloxy" enzymatic solution by mixing 60 &mu;l water, 20 &mu;l of 5X buffer A, 20 &mu;l of catalase, 1 mg glucose oxidase and 100 &mu;l of 10 mg/ml BSA. Filter the solution using a 0.22 &mu;m centrifuge filter.</li>
 <li>Gently mix the buffer from Step 2 with the "gloxy" solution from Step 3 avoiding the formation of air bubbles.</li>
 <li>Flow the IB through the channel at a constant rate of 5 &mu;l/min using a mechanical syringe pump to control the flow rate. Bubble argon gas into the IB reservoir to prevent atmospheric oxygen from re-dissolving into the buffer.</li>
</ol>
The surface scanning, molecule localization and acquisition of single molecule intensity traces is controlled by a LabView program that permits automated data collection1. The program scans 20x20 &mu;m2 areas at 0.2 &mu;m resolution, with a rate of 0.1 &mu;m/ms. Data is immediately processed to yield intensity-weighted locations of all pixels above the background counts. Once the immobilized molecules are found, they are moved one by one into the confocal volume and the intensities of the donor and acceptor fluorophore are recorded as a function of time until both fluorophores photobleach. The stage is then programmed to move to a new origin 100 &mu;m away, and the scanning process is repeated.
4 - Representative Results
Below are some images showing the assembled microfluidic cell before surface activation and after being mounted on the microscope ready for molecule immobilization. If the channel is activated successfully, surface scans should be similar to the scans depicted below showing emission of the donor dye (green) and acceptor (red) after direct donor excitation. These molecules are moved one by one into the probing volume and the fluorescence is recorded over time until both dyes photobleach, as shown in the single molecule traces. From traces like these one can obtain the FRET efficiency which informs on the instantaneous donor-acceptor separation, which in turn is used to explore the structure and dynamics of molecules of biological interest.
<img src="/files/ftp_upload/1542/1542fig1tmb.jpg" alt="Figure 1" border="0" /> Figure 1: Microfluidic cell with four channels, each with inlet/outlet capillaries. Please <a href="https://www.jove.com/files/ftp_upload/1542/1542fig1.jpg">click here</a> to see a larger version of figure 1.
<img src="/files/ftp_upload/1542/1542fig2tmb.jpg" alt="Figure 2" border="0" /> Figure 2: Cell mounted on the microscope ready for measurements. Please <a href="https://www.jove.com/files/ftp_upload/1542/1542fig2.jpg">click here</a> to see a larger version of figure 2.
<img src="/files/ftp_upload/1542/1542fig3tmb.jpg" alt="Figure 3" border="0" /> Figure 3: Setup for sm-FRET measurements. Please <a href="https://www.jove.com/files/ftp_upload/1542/1542fig3.jpg">click here</a> to see a larger version of figure 3.
<img src="/files/ftp_upload/1542/1542fig4tmb.jpg" alt="Figure 4" border="0" /><img src="/files/ftp_upload/1542/1542fig5tmb.jpg" alt="Figure 5" border="0" /> Figures 4 and 5: Red and green channel of a surface scan of immobilized FRET molecules excited with green laser. Please click to see a larger version of <a href="https://www.jove.com/files/ftp_upload/1542/1542fig4.jpg">figure 4</a>, or <a href="https://www.jove.com/files/ftp_upload/1542/1542fig5.jpg">figure 5</a>.
<img src="/files/ftp_upload/1542/1542fig6tmb.jpg" alt="Figure 6" border="0" /><img src="/files/ftp_upload/1542/1542fig7tmb.jpg" alt="Figure 7" border="0" /> <img src="/files/ftp_upload/1542/1542fig8tmb.jpg" alt="Figure 8" border="0" /><img src="/files/ftp_upload/1542/1542fig9tmb.jpg" alt="Figure 9" border="0" /> Figures 6, 7, 8, and 9: Intensity trajectories of donor (green) and acceptor (red) fluorophores labeled to a DNA molecule 8, 10, 16 and 18 basepairs apart. Please click to see a larger version of 
<a href="https://www.jove.com/files/ftp_upload/1542/1542fig6.jpg">figure 6</a>, <a href="https://www.jove.com/files/ftp_upload/1542/1542fig7.jpg">figure 7</a>,<a href="https://www.jove.com/files/ftp_upload/1542/1542fig8.jpg">figure 8</a>, or <a href="https://www.jove.com/files/ftp_upload/1542/1542fig9.jpg">figure 9</a>.

<ol>
	<li>Selvin, P. R., Ha, T. <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+Techniques,+a+Laboratory+Manual.">Single-Molecule Techniques, a Laboratory Manual.</a> CSHL Press. , (2008).</li><li>Sabanayagam, C. R., Eid, J. S., Meller, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+scanning+confocal+microscope+for+single+molecule+analysis.">High-throughput scanning confocal microscope for single molecule analysis.</a> Applied Pys. Letters. , 84-87 (2004).</li><li>Sabanayagam, C. R., Eid, J. S., Meller, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+Fluorescence+resonance+energy+transfer+to+measure+distances+along+individual+DNA+molecules:+Corrections+due+to+nonideal+transfer.">Using Fluorescence resonance energy transfer to measure distances along individual DNA molecules: Corrections due to nonideal transfer.</a> J. Chem. Phys. 122, 61103-61105 (2004).</li><li>Echeverra, A., Aitken, C., Marshall, R. A., Puglisi, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+Oxygen+Scavenging+System+for+Improvement+of+Dye+Stability+in+Single-Molecule+Fluorescence+Experiments.">An Oxygen Scavenging System for Improvement of Dye Stability in Single-Molecule Fluorescence Experiments.</a> Biophysical Journal. 94, 1826-1835 (2008).</li><li>Rasnik, I., McKinney, S. A., Ha, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonblinking+and+long-lasting+single+molecule+fluorescence+imaging.">Nonblinking and long-lasting single molecule fluorescence imaging.</a> Nature Methods. 3 (11), 891-893 (2006).</li><li>Boukobza, E., Sonnenfeld, A., Haran, G., Puglisi, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immobilization+in+Surface-Tethered+Lipid+Vesicles+as+a+New+Tool+for+Single+Biomolecule+Spectroscopy.">Immobilization in Surface-Tethered Lipid Vesicles as a New Tool for Single Biomolecule Spectroscopy.</a> J. Phys. Chem. 105, 12165-12170 (2001).</li><li>Di Fiori, N., Meller, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Article in preparation. , .</li></ol>

Although the above protocol has been optimized for our particular system and for a specific choice of dyes (TMR and ATTO647N), it is by no means the only proved method. Recently, an alternative oxygen scavenging system consisting of protocatechuic acid (PCA)/protocatechuate-3,4-dioxygenase (PCD) was shown to increase the photostability of certain dyes4. Likewise, other triplet-state quenchers such as &beta;-mercaptoethanol (BME) can be used instead of Trolox, although these might not suppress long-lasting blinking of some dyes, in particular Cy55.
Although immobilization via biotin-streptavidin-biotinylated bovine serum albumin (BSA) is the preferred choice for nucleic acids studies, a polyethylene glycol (PEG)-coated surface is better suited for studies of proteins since it prevents non-specific adhesion1. Additionally, sub-diffraction-limited size vesicles can be used to encapsulate the biomolecule of interest in cases where it can be affected by surface interactions6.
In this article we have used a confocal setup equipped with silicone avalanche photodiodes to acquire the fluorescence intensity traces. This protocol works equally well with Total Internal Reflection Fluorescence microscopes (TIRF) equipped with CCD cameras. Regardless of the microscopy used, sm-FRET can inform on spatial resolutions approaching the angstrom when the donor and acceptor signal are properly corrected7.

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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<tr>
<td>Fused silica capillary tubing</td>
<td>&nbsp;</td>
<td>Polymicro Technologies</td>
<td>TSP150375</td>
<td>&nbsp;</td>
<tr>
<td>Fused silica cover slip</td>
<td>&nbsp;</td>
<td>Esco Products</td>
<td>R425000</td>
<td>&nbsp;</td>
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<td>Tubing</td>
<td>&nbsp;</td>
<td>Diba Industries</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Biotinylated BSA</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>A8549</td>
<td>&nbsp;</td>
<tr>
<td>BSA</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>A9085</td>
<td>&nbsp;</td>
<tr>
<td>Streptavidin</td>
<td>&nbsp;</td>
<td>Thermo Scientific</td>
<td>0021122</td>
<td>&nbsp;</td>
<tr>
<td>Sylgard 184 Elastomer Kit</td>
<td>&nbsp;</td>
<td>DowCorning</td>
<td>3097366-1004</td>
<td>Silicone Elastomer Kit</td>
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<td>Trolox</td>
<td>&nbsp;</td>
<td>Aldrich</td>
<td>238813</td>
<td>&nbsp;</td>
<tr>
<td>Catalase</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>C3155</td>
<td>&nbsp;</td>
<tr>
<td>Glucose Oxidase</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>G2133</td>
<td>&nbsp;</td>
<tr>
<td>D-Glucose</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>G7528</td>
<td>&nbsp;</td>
<tr>
<td>Kwik-Cast Sealant</td>
<td>&nbsp;</td>
<td>World Precision Instruments</td>
<td>Kwik-Cast</td>
<td>Silicone Casting Compound</td>
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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.

Watch this Scientific Journal Video about Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules at JoVE.com

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

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, ...

automated-system-for-single-molecule-fluorescence-measurements

Research

JoVE Journal

Biology

Single Molecule Methods for Monitoring Changes in Bilayer Elastic Properties

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein interactions and more general lipid bilayer-protein interactions. These interactions are particularly important in pharmacological research, as many current pharmaceuticals on the market can alter the lipid bilayer material properties, which can lead to altered membrane protein function.   The formation of gramicidin channels are dependent on conformational changes in gramicidin subunits which are in turn dependent on the properties of the lipid.  Hence the gramicidin channel  current is a reporter of altered properties of the bilayer due to certain compounds.  

Membrane protein function is regulated by the cell membrane lipid composition. This video-article details how to form a patch using bilayer patch electrodes, as well as how to use gramicidin channels as reporters of altered membrane properties.

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein ...

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.


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 ...

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 ...

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 ...

Open Source High Content Analysis Utilizing Automated Fluorescence Lifetime Imaging Microscopy

We present an open source high content analysis instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions using F&#246;rster resonance energy transfer (FRET) based readouts of fixed or live cells in multiwell plates. This provides a means to screen for cell signaling processes read out using intramolecular FRET biosensors or intermolecular FRET of protein interactions such as oligomerization or heterodimerization, which can be used to identify binding partners. We describe here the functionality of this automated multiwell plate FLIM instrumentation and present exemplar data from our studies of HIV Gag protein oligomerization and a time course of a FRET biosensor in live cells. A detailed description of the practical implementation is then provided with reference to a list of hardware components and a description of the open source data acquisition software written in &#181;Manager. The application of FLIMfit, an open source MATLAB-based client for the OMERO platform, to analyze arrays of multiwell plate FLIM data is also presented. The protocols for imaging fixed and live cells are outlined and a demonstration of an automated multiwell plate FLIM experiment using cells expressing fluorescent protein-based FRET constructs is presented. This is complemented by a walk-through of the data analysis for this specific FLIM FRET data set.

We present an open source high content analysis (HCA) instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions using F&#246;rster resonance energy transfer (FRET) based readouts. Data acquisition for this openFLIM-HCA instrument is controlled by software written in &#181;Manager and data analysis is undertaken in FLIMfit.

We present an open source high content analysis instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions ...

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH allows the measurement of cell-to-cell variability in mRNA copy number of genes of interest, as well as the subcellular location of the transcripts. The main steps involved are fixation of the bacterial cell culture, permeabilization of cell membranes, and hybridization of the target transcripts with sets of commercially available short fluorescently-labeled oligonucleotide probes. smFISH can allow the imaging of the transcripts of multiple genes in the same cell, with limitations imposed by the spectral overlap between different fluorescent markers. Following completion of the protocol illustrated below, cells can be readily imaged using a microscope coupled with a camera suitable for low-intensity fluorescence. These images, together with cell contours obtained from segmentation of phase contrast frames, or from cell membrane staining, allow the calculation of the mRNA copy number distribution of a sample of cells using open-source or custom-written software. The labeling method described here can also be applied to image transcripts with stochastic optical reconstruction microscopy (STORM).

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

This manuscript describes a method for labeling individual messenger RNA (mRNA) transcripts with fluorescently-labeled DNA probes, for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH is a visualization method that allows the simultaneous detection, localization, and quantification of single mRNA molecules in fixed individual cells.

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ ...

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 ...

Reconstruction of Single-Cell Innate Fluorescence Signatures by Confocal Microscopy

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing the innate cellular fluorescence signature from each individual live cell in a population distributed in a three-dimensional (3D) space. The innate fluorescence signature of a cell is a collection of fluorescence signals emitted by various biomolecules within the cell. Previous studies established that innate fluorescence signatures reflect various cellular properties and differences in physiological status and are a rich source of information for cell characterization and identification. Innate fluorescence signatures have been traditionally analyzed at the population level, necessitating a clonal culture, but not at the single-cell level. CRIF is particularly suitable for studies that require 3D resolution and/or selective extraction of fluorescence signals from individual cells. Because the fluorescence signature is an innate property of a cell, CRIF is also suitable for tag-free prediction of the type and/or physiological status of intact and single cells. This method may be a powerful tool for streamlined cell analysis, where the phenotype of each single cell in a heterogenous population can be directly assessed by its autofluorescence signature under a microscope without cell tagging.

Here, a protocol is presented for optically extracting and cataloging innate cellular fluorescence signatures (i.e., cellular autofluorescence) from every individual live cell distributed in a three-dimensional space. This method is suitable for studying the innate fluorescence signature of diverse biological systems at a single-cell resolution, including cells from bacteria, fungi, yeasts, plants, and animals.

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing ...

Parallel High Throughput Single Molecule Kinetic Assay for Site-Specific DNA Cleavage

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable of measuring SSDC in many single DNA molecules simultaneously. Bead-tethered substrate DNAs, each containing a single copy of the target sequence, are prepared in a microfluidic flow channel. An external magnet applies a weak force to the paramagnetic beads. The integrity of up to 1,000 individual DNAs can be monitored by visualizing the microbeads under darkfield imaging using a wide-field, low magnification objective. Injecting of a restriction endonuclease, NdeI, initiates the cleavage reaction. Video microscopy is used to record the exact moment of each DNA cleavage by observing the frame in which the associated bead moves up and out of the focal plane of the objective. Frame-by-frame bead counting quantifies the reaction, and an exponential fit determines the reaction rate. This method allows collection of quantitative and statistically significant data on single molecule SSDC reactions in a single experiment.

A highly parallel method for measuring the site-specific cleavage of DNA at the single molecule level is described. This protocol demonstrates the technique using the restriction endonuclease NdeI. The method can easily be modified to study any process that results in site-specific DNA cleavage.

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable ...