The authors would like to thank Anke R&uuml;ttgeroth and Karina Zimmermann for technical assistance. This work was supported by the Deutsche Forschungsgemeinschaft (grant NI 1301/1-1 to V.O.N) and University of G&ouml;ttingen Medical Center ("pro futura" grant to V.O.N.).

1. Setting up a FRET Imaging Microscope
In principle, any inverted fluorescence microscope which is available in the lab and has a camera port can be adapted for FRET imaging. The final setup should include the following crucial components: a microscope, a light source with or without additional shutter, a beam-splitter for emission light and a CCD-camera (see Figure 1). The hardware devices, especially the light source, the shutter and the camera are integrated into and controlled by the imaging software which allows image acquisition and analysis. Below we describe a procedure to assemble a simple FRET system from commercially available components.
<ol>
 <li>Connect your light source to the microscope. For example, use a single-wavelength light emitting diode (CoolLED pE-100, 440 nm) which selectively excites enhanced cyan fluorescent protein (CFP) used as a donor in most FRET biosensors. It can be directly and easily connected to the epifluorescence illumination port of the microscope. There are also multi-wavelength LEDs available which can be connected in a similar way. Instead of LED, other standard light sources such as XBO75 xenon arc lamp (often used for Olympus and Zeiss microscopes) or HBO mercury lamp (usually installed on Nikon microscopes) can be used. In the case of a fluorescent lamp, you should also place a shutter between the lamp and illumination port to enable software-assisted control of the excitation light coming onto your sample. CoolLED does not require any additional shutter since it can be directly switched on and off by the software. The disadvantage of the single-color LED systems is a limited number of excitation wavelengths, while a fluorescent lamp with various filter sets is a more flexible option, especially when working with multiple and bimolecular biosensors. Alternatively, monochromator light sources (e.g. Polychrome V, TILLPhotonics) can be used; they usually have an integrated shutter which can be software-controlled via a trigger signal.</li>
 <li>Place an appropriate filter cube into the microscope. For routine FRET measurements with CFP and enhanced yellow fluorescent protein (YFP) or any of their variants as a FRET pair, we use a simple filter cube containing an ET436/30M excitation filter (which can be omitted when LED is used, but is indispensable when using a xenon or mercury lamp rather than LED) and a DCLP455 dichroic mirror. The microscope should also be equipped with an objective suitable for a good-resolution fluorescence microscopy, for example with a plan-fluor, plan-neofluar or plan-apochromat 40x, 60x or 100x oil-immersion objective.</li>
 <li>Switch on the fluorescent light and check whether the light spot is evenly distributed across the field of view. If this is not the case, additional alignment of the LED or the lamp is required to achieve the optimal illumination of the specimen. This can be performed by using the screws which position the diode or the lamp in space.</li>
 <li>Connect the beam-splitter via a C-mount to one of the microscope's emission ports. For example, use the DV2 DualView (Photometrics) which splits the emission light into two (donor and acceptor) channels which can be simultaneously monitored on a single CCD camera chip. Alternatively, there are other comparable products such as Optosplit (Cairn Research) or a beam splitter integrated into the Hamamatsu ORCA-D2 dual-wavelength camera. For the CFP/YFP FRET pair, we use the 05-EM filter set containing the 505dcxr dichroic mirror plus ET480/30M and ET535/40M emission filters for CFP and YFP, respectively, which is supplied with the DV2. Instead of a beam-splitter, two filter cubes for the donor (containing the ET436/30M excitation filter for CFP, the DCLP455 dichroic mirror and the CFP emission filter) and acceptor (containing the CFP excitation filter, DCLP455 and the YFP emission filter) channels are used in many FRET systems. Alternatively, one filter cube without any emission filter and an automatic emission filter wheel position before the camera can be installed. In this case, a motorized microscope or the filter wheel alternates between the two emission filter positions within ~200-300 msec to perform the ratiometric imaging. This minor delay is acceptable when imaging rather slow intracellular processes, such as cAMP signals, where truly simultaneous acquisition of both channels is not critical.</li>
 <li>Connect the CCD camera (use for example ORCA-03G or ORCA-R2 from Hamamatsu Photonics) to the beam splitter. Use a FireWire cable to connect the camera to the IEEE1394 computer interface, as described in the manual supplied with the camera. Install camera drivers without switching on the camera.</li>
 <li>Finally, to establish the communication between the computer and the light source, connect the Arduino I/O board (use for example Arduino Duemilanove or Arduino Uno) to the LED or to the shutter by using a BNC cable which should contain a normal BNC plug on the LED side and two single wires on the other end connected into the pins GND (0) and 8 of the board as shown in Figure 2. The assembled board can be directly connected to a USB-port of your computer.</li>
</ol>
2. Setting up the Imaging Software
To control and synchronize the light source with image capturing by the camera, imaging software should be installed on the computer. There are several commercially available software packages including MetaFluor (Molecular Devices), Slidebook (Intelligent Imaging Innovations), VisiView (Visitron Systems). Here we demonstrate the use of the open-source Micro-Manager freeware which offers a high degree of flexibility for low-cost imaging.
<ol>
 <li>Download this software at <a href="http://valelab.ucsf.edu/~MM/MMwiki/index.php/Micro-Manager_Version_Archive">http://valelab.ucsf.edu/~MM/MMwiki/index.php/Micro-Manager_Version_Archive</a>. We recommend installing the 1.4.5 release which is easily configurable in our hands.</li>
 <li>Connect the Arduino board to a USB port of your computer. Download the software to control the Arduino board from <a href="http://www.arduino.cc/en/Main/software" target="_blank">http://www.arduino.cc/en/Main/software</a>. Follow the instructions found on this website and run this software just one time prior to starting Micro-Manager. Upload a code for use of the board with Micro-Manager software. The code can be downloaded at <a href="http://valelab.ucsf.edu/~MM/MMwiki/index.php/Arduino" target="_blank">http://valelab.ucsf.edu/~MM/MMwiki/index.php/Arduino</a>.</li>
 <li>Switch on the LED and the camera. Start the software and configure Micro-Manager communication with the camera and LED (other light source and shutter) by selecting Tools&gt;Hardware Configuration Wizard (see Figure 3). Add the required components including your camera (i.e. Hamamatsu_ DCAM) and Arduino Board (add Arduino-Switch, Arduino-Hub and Arduino-Shutter devices). During the next steps, use the default settings suggested by the wizard. Save the new system configuration when prompted by the wizard.</li>
 <li>Use the main menu to open Tools&gt;Device Property Browser. Scroll down to "Arduino Switch State" and select "1". Close the dialog. Make sure that the "auto-shutter" box is ticked in the main software dialog. Press File&gt;Save System State to save the software configuration which should be opened anytime after starting the software. This will establish the communication between the board and the software needed to perform image acquisition.</li>
 <li>Press "Live" button to monitor the signal coming from the camera. Make sure that fluorescent light goes on any time "Live" or "Snap" function is selected. Before starting the first measurements, follow the instructions supplied with your beam splitter to perform the optical alignment of both channels.</li>
</ol>
3. Cell Culture and Transfections
<ol>
 <li>Prepare 6-well plates with autoclaved round 24 mm glass coverslides (1 coverslide per well). Alternatively, glass-bottomed cell-culture dishes can be used.</li>
 <li>Plate 293A cells onto the plates or dishes in D-MEM medium (supplemented with 10% FCS, 1% L-glutamine and 1% penicillin/streptomycin solution) so that the cells reach 50-70% confluency after one day.</li>
 <li>24 hr after plating, transfect the cells in a laminar flow bench with a FRET sensor plasmid using the calcium phosphate transfection method (see 3.5). cAMP biosensor plasmids can be obtained from our group upon request. The reader is also referred to the comprehensive reviews7,8 describing other available cAMP biosensors.</li>
 <li>Before transfecting cells for the first time, prepare transfection reagents. Make up a 2.5 M CaCl2 and 2xBBS solutions (the latter containing 1.5 mM Na2HPO4, 50 mM BES, 280 mM NaCl, adjust to pH 6.95 with NaOH) in deionized water. Sterile filtrate the solutions using a regular 0.2 &mu;m filter.</li>
 <li>To transfect a 6-well plate or 6 glass-bottomed dishes, mix 10 &mu;g of FRET sensor plasmid, 50 &mu;l of 2.5M CaCl2 solution and sterile water up to 500 &mu;l. Mix well.</li>
 <li>Add 500 &mu;l of 2x BBS. Mix well and incubate the mixture for 10 min at room temperature.</li>
 <li>Pipette 165 &mu;l of the transfection mix dropwise onto each well or dish. Gently stir the plate and put it back into the incubator. Cells are usually ready for FRET measurements 24 hr after transfection. Make sure that cells have not reached confluency at this point, as this might influence the activity of several cell-surface receptors.</li>
</ol>
4. FRET Measurements in Live Cells
<ol>
 <li>Before measuring for the first time, prepare the FRET buffer containing 144 mM NaCl, 5.4 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 10 mM HEPES in deionized water and adjust the pH to 7.3 with NaOH. Dilute the compounds to be used in your imaging experiments with the FRET buffer.</li>
 <li>Start the imaging software. Load previously defined system configuration. Select File&gt;Load System State to choose the previously configured system state.</li>
 <li>Mount a coverslide with adherent transfected 293A cells in the imaging chamber (e.g. Attofluor cell chamber). Wash the cells once with FRET buffer and add 400 &mu;l of FRET buffer. When using glass-bottomed cell-culture dishes wash the adherent cells and add 2 ml of the buffer per dish. We perform all measurements at room temperature in the FRET buffer containing HEPES, so that no CO2 control is necessary.</li>
 <li>Put some immersion oil onto the objective and transfer the imaging chamber onto the microscope. Focus on the cell layer using transillumination light.</li>
 <li>Switch on the fluorescent light by pressing the "Live" button, and select a cell for the experiment. Choose a cell with an optimal sensor expression, meaning that it should be not too bright and not too dim. After finding an appropriate cell, switch off the fluorescent light immediately to avoid photobleaching of the FRET sensor.</li>
 <li>Adjust the exposure time under the "Camera settings" (usually 10-50 msec) in a way that leads to a good signal-to-noise ratio of the acquired image after pressing the "Snap" button. Too long excitation times might result in photobleaching, while too short times result in low image quality.</li>
 <li>Press the "Multi-D Acq." button and set the number of time points and the time interval for image acquisition. For our cAMP-FRET sensor, we acquire one image every 5 s.</li>
 <li>Start the measurements by pressing the "Acquire" button.</li>
 <li>During any measurement, one can use the "FRET online" plug-in (available in the Online Supplement, all plug-ins must be copied into the "plugins" folder of your Micro-Manager software before starting it) to monitor FRET ratio changes online. Run this plug-in and select a region of interest in the FRET ratio image using the "Freehand selections" tool. Add the region into the ROI manager and press the "Get average" button in the "Time series analyzer" window. The FRET ratio trace will be displayed. To update the trace during the measurement, run the "FRETratioOnline2" plug-in and press the "GetAverage" button.</li>
 <li>As soon as the FRET ratio has reached a stable baseline apply the desired compound by accurately pipetting it into the dish/chamber. To treat cells with pharmacological compounds, a perfusion system instead of simple pipetting can be used at this stage.</li>
 <li>After finishing the experiment, save the time-lapse stack of images. Remove the measurement chamber from the microscope and clean the objective using an objective tissue.</li>
 <li>Go back to step 4.3 to repeat the measurement with a new sample.</li>
</ol>
5. Offline Data Analysis
FRET imaging data can be analyzed offline at any time after the experiment using ImageJ software. As a supplement to this protocol, we provide the "FREToffline" plug-in used in our laboratory to split the acquired images into donor and acceptor channels and to measure fluorescent intensities in multiple regions of interest. These intensities can further be copy-pasted into an Excel or Origin spreadsheet to calculate the corrected FRET ratio. To visualize the FRET changes of unimolecular biosensors, simple ratiometry is often used. In this case, only the donor fluorophore (CFP) is excited, and two images are taken at CFP and YFP emission peaks. The calculated YFP/CFP ratio (sometimes also referred to as FRET/CFP ratio) represents the degree of FRET between the two fluorophores. In unimolecular biosensors, the numbers of CFP and YFP moieties are equal, so that the simple ratiometry is sufficient to represent the FRET efficiency12.
<ol>
 <li>Use ImageJ software to open the experiment file by selecting Plugins&gt;Micro-Manager&gt;Open Micro-Manager File.</li>
 <li>Run the "FREToffline" plug-in which splits the time-lapse stack into individual CFP and YFP channels.</li>
 <li>If background correction is required, this can be performed using ImageJ software.</li>
 <li>Click on the YFP stack of images and select one or several regions of interest using the "Freehand selections" tool and add them to the "MultiMeasure" plug-in window by pressing the "Add" button.</li>
 <li>Select the regions of interests in the "MultiMeasure" window and press "Multi" to obtain a table with mean grey values for each frame and each region. Copy the data into clipboard by selecting all using Ctrl+A and by pressing Ctrl+C. Open an Excel or Origin spreadsheet and paste the data by pressing Ctrl+V. 
 The measurements performed by the program can be configured under Analyze&gt;Set Measurements where you can define the parameters to be measured. We usually select only "Mean grey value" in this dialog.</li>
 <li>Click onto the CFP stack of images. Perform the same as described in 5.5. Paste the CFP intensity data into the same Excel or Origin spreadsheet.</li>
 <li>Using Excel or Origin software, calculate the corrected FRET ratio. When simple single-chain unimolecular biosensors are imaged, we correct only for the bleedthrough of the donor fluorescence into the acceptor channel. In this case, the corrected acceptor/donor ratio is: 
 Ratio = (YFP - B x CFP) / CFP 
 Where B is the correction factor which can be determined by transfecting cells with CFP plasmid and measuring a percentage of the donor fluorescence in the YFP channel (B =YFP/CFP). Omitting this correction for unimolecular biosensors is possible, since it would only affect the overall amplitude of the FRET response without any qualitative effect on the shape of the curve.</li>
</ol>
6. Representative Results
Figure 1 shows an example of a fully assembled FRET imaging setup consisting of a Nikon inverted microscope, CoolLED, DV2 DualView and the Hamamatsu ORCA-03G CCD camera. To establish the communication between the hardware components and the computer, the I/O Arduino board is connected to the computer and to the CoolLED as shown in Figure 2. To control the light source and image capturing by the camera in a synchronized fashion, Micro-Manager software has to be installed and properly configured (see Figure 3). This freeware can be easily adapted to individual experimental needs by adding necessary plug-ins. Figure 4A shows a representative raw FRET ratio trace from a measurement using the cAMP sensor Epac1-camps13 expressed in 293A cells to monitor the effects of &beta;-adrenergic agonist isoproterenol applied at time point 150 sec and the &beta;-blocker propranolol added at time point 300 sec (performed as described in 4.3-4.10). These data can be analyzed offline and corrected for the bleedthrough of CFP into the YFP channel as described in 5.1-5.7 to obtain the corrected FRET ratio trace shown in Figure 4B. This representative experiment shows a slow decrease in the monitored FRET ratio upon isoproterenol treatment which indicates an increase of intracellular cAMP. Propranolol as a &beta;-blocker reverses the isoproterenol signal, leading to a decrease of cAMP to basal levels. These changes in FRET signal can be monitored online during any experiments (as described in 4.9). Such experiments can be performed with a variety of commonly used biosensors designed to monitor different second messengers or biochemical processes.
<img alt="Figure 1" src="/files/ftp_upload/4081/4081fig1.jpg" /> 
Figure 1. Layout of the FRET imaging setup comprised of a CoolLED, inverted Nikon microscope, DV2 DualView, and ORCA-03G CCD camera.
<img alt="Figure 2" src="/files/ftp_upload/4081/4081fig2.jpg" /> 
Figure 2. Arduino I/O board and its connections. The board is positioned in a self-mounted plastic box. A standard BNC cable connects the LED to the pins 8 and GND (0) of the board.
<img alt="Figure 3" src="/files/ftp_upload/4081/4081fig3.jpg" /> 
Figure 3. Screenshots demonstrating integration of the system components using Hardware Configuration Wizard. A) Start the Hardware Configuration Wizard. B) Add the required devices as mentioned in 2.3. <a href="https://www.jove.com/files/ftp_upload/4081/4081fig3large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 4" src="/files/ftp_upload/4081/4081fig4.jpg" /> 
Figure 4. A representative FRET experiment which measures cAMP levels in 293A cells transfected with Epac1-camps. First, cells were stimulated with the &beta;-adrenergic agonist isoproterenol (100 nM, at frame 30 or at 150 sec) to increase cAMP (observed as a decrease in YFP/CFP FRET ratio). Cells were subsequently treated with the &beta;-blocker propranolol (10 &mu;M, at frame 60 or at 300 sec) which leads to an increase of FRET ratio, reflecting a decrease in cAMP. A) Raw online FRET ratio trace from one region of interest corresponding to a single cell monitored during the experiment. B) Corrected ratio trace after offline data analysis performed as described in 5.1-5.7.

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No conflicts of interest declared.

in this protocol, we demonstrate how to build a simple low-cost but powerful FRET imaging system for routine applications with a variety of available biosensors. The system presented here is designed for CFP and YFP, or similar types of fluorescent proteins, as the donor-acceptor pair. Meanwhile, other individual biosensors become available which use for example green and red fluorescent proteins14. To adapt the described system for other colors, appropriate light sources and/or filter sets should be selected....

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent/equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>BES Buffer Grade</td>
 <td>AppliChem</td>
 <td>A1062</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CaCl2 dihydrate</td>
 <td>Sigma-Aldrich</td>
 <td>C5010 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Glass coverslides</td>
 <td>Thermo Scientific</td>
 <td>004710781</td>
 <td>Diameter 24 mm</td>
 </tr>
 <tr>
 <td>Glass-bottomed cell-culture dishes</td>
 <td>World Precision Instruments</td>
 <td>FD3510-100 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>D-MEM medium</td>
 <td>Biochrom AG</td>
 <td>F0445</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fetal calf serum (FCS)</td>
 <td>Thermo Scientific</td>
 <td>SH30073.02</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>L-Glutamine</td>
 <td>Biochrom AG</td>
 <td>K0283</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Sigma</td>
 <td>H4034</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>Sigma</td>
 <td>P5405</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>MgCl2 hexahydrate</td>
 <td>AppliChem</td>
 <td>A4425</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>AppliChem</td>
 <td>A1149</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Na2HPO4</td>
 <td>Sigma-Aldrich</td>
 <td>S9707</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Penicillin/Streptomycin</td>
 <td>Biochrom AG</td>
 <td>A2213</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Inverted fluorescent microscope</td>
 <td>e.g. Nikon</td>
 <td>Request at Nikon</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CoolLED</td>
 <td>CoolLED</td>
 <td>pE-100 </td>
 <td>440 nm</td>
 </tr>
 <tr>
 <td>DualView</td>
 <td>Photometrics</td>
 <td>DV2-SYS</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>DualView filter slider</td>
 <td>Photometrics</td>
 <td>05-EM</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CFP/YFP filter set</td>
 <td>Chroma Technology</td>
 <td>49052</td>
 <td>without the emission filter</td>
 </tr>
 <tr>
 <td>ORCA-03G camera</td>
 <td>Hamamatsu Photonics</td>
 <td>C8484-03G02 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Arduino I/O board </td>
 <td>Sparkfun Electronics</td>
 <td>DEV-00666</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Attofluor cell chamber </td>
 <td>Invitrogen</td>
 <td>A-7816</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Personal computer with WindowsXP or Windows7 system </td>
 <td>Any supplier</td>
 <td>&nbsp;</td>
 <td>Include hard-drive with high capacity</td>
 </tr>
 </tbody>
</table>

fret microscopy for real-time monitoring of signaling events in live cells using unimolecular biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

Watch this Scientific Journal Video about FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors at JoVE.com

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

fret-microscopy-for-real-time-monitoring-signaling-events-live-cells

Research

JoVE Journal

Biology

23.0K Views.  Georg August University Medical Center, Göttingen, Germany. Förster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

Video: FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

Combining QD-FRET and Microfluidics to Monitor DNA Nanocomplex Self-Assembly in Real-Time

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically functional inside the cell, nucleic acid-based therapeutics require an efficient intracellular delivery system. One widely adopted approach is to complex DNA with a gene carrier to form nanocomplexes via electrostatic self-assembly, facilitating cellular uptake of DNA while protecting it against degradation. The challenge lies in the rational design of efficient gene carriers, since premature dissociation or overly stable binding would be detrimental to the cellular uptake and therapeutic efficacy. Nanocomplexes synthesized by bulk mixing showed a diverse range of intracellular unpacking and trafficking behavior, which was attributed to the heterogeneity in size and stability of nanocomplexes. Such heterogeneity hinders the accurate assessment of the self-assembly kinetics and adds to the difficulty in correlating their physical properties to transfection efficiencies or bioactivities. We present a novel convergence of nanophotonics (i.e. QD-FRET) and microfluidics to characterize the real-time kinetics of the nanocomplex self-assembly under laminar flow. QD-FRET provides a highly sensitive indication of the onset of molecular interactions and quantitative measure throughout the synthesis process, whereas microfluidics offers a well-controlled microenvironment to spatially analyze the process with high temporal resolution (~milliseconds). For the model system of polymeric nanocomplexes, two distinct stages in the self-assembly process were captured by this analytic platform. The kinetic aspect of the self-assembly process obtained at the microscale would be particularly valuable for microreactor-based reactions which are relevant to many micro- and nano-scale applications. Further, nanocomplexes may be customized through proper design of microfludic devices, and the resulting QD-FRET polymeric DNA nanocomplexes could be readily applied for establishing structure-function relationships. 

We present a novel and powerful integration of nanophotonics (QD-FRET) and microfluidics to investigate the formation of polyelectrolyte polyplexes, which is expected to provide better control and synthesis of uniform and customizable polyplexes for future nucleic acid-based therapeutics.

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically ...

Real-time Monitoring of Ligand-receptor Interactions with Fluorescence Resonance Energy Transfer

FRET is a process whereby energy is non-radiatively transferred from an excited donor molecule to a ground-state acceptor molecule through long-range dipole-dipole interactions1. In the present sensing assay, we utilize an interesting property of PDA: blue-shift in the UV-Vis electronic absorption spectrum of PDA (Figure 1) after an analyte interacts with receptors attached to PDA2,3,4,7. This shift in the PDA absorption spectrum provides changes in the spectral overlap (J) between PDA (acceptor) and rhodamine (donor) that leads to changes in the FRET efficiency. Thus, the interactions between analyte (ligand) and receptors are detected through FRET between donor fluorophores and PDA. In particular, we show the sensing of a model protein molecule streptavidin. We also demonstrate the covalent-binding of bovine serum albumin (BSA) to the liposome surface with FRET mechanism. These interactions between the bilayer liposomes and protein molecules can be sensed in real-time. The proposed method is a general method for sensing small chemical and large biochemical molecules. Since fluorescence is intrinsically more sensitive than colorimetry, the detection limit of the assay can be in sub-nanomolar range or lower8. Further, PDA can act as a universal acceptor in FRET, which means that multiple sensors can be developed with PDA (acceptor) functionalized with donors and different receptors attached on the surface of PDA liposomes.

We demonstrate FRET between conjugated polymer polydiacetylene (PDA) and fluorophore attached to the surface of PDA liposomes for the sensing of biomolecules. PDA liposomes also contained receptor molecules on their surfaces for biomolecules to be used as probes. Ligand-receptor interactions lead to changes in the FRET efficiency between the fluorophore and PDA which is the basis of the sensing mechanism.

FRET is a process  whereby energy is non-radiatively transferred from an excited donor molecule to  a ground-state acceptor molecule through long-range ...

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) ...

Autonomously Bioluminescent Mammalian Cells for Continuous and Real-time Monitoring of Cytotoxicity

Mammalian cell-based in vitro assays have been widely employed as alternatives to animal testing for toxicological studies but have been limited due to the high monetary and time costs of parallel sample preparation that are necessitated due to the destructive nature of firefly luciferase-based screening methods. This video describes the utilization of autonomously bioluminescent mammalian cells, which do not require the destructive addition of a luciferin substrate, as an inexpensive and facile method for monitoring the cytotoxic effects of a compound of interest. Mammalian cells stably expressing the full bacterial bioluminescence (luxCDABEfrp) gene cassette autonomously produce an optical signal that peaks at 490 nm without the addition of an expensive and possibly interfering luciferin substrate, excitation by an external energy source, or destruction of the sample that is traditionally performed during optical imaging procedures. This independence from external stimulation places the burden for maintaining the bioluminescent reaction solely on the cell, meaning that the resultant signal is only detected during active metabolism. This characteristic makes the lux-expressing cell line an excellent candidate for use as a biosentinel against cytotoxic effects because changes in bioluminescent production are indicative of adverse effects on cellular growth and metabolism. Similarly, the autonomous nature and lack of required sample destruction permits repeated imaging of the same sample in real-time throughout the period of toxicant exposure and can be performed across multiple samples using existing imaging equipment in an automated fashion.

Mammalian cells expressing the bacterial bioluminescence gene cassette (lux) produce light autonomously. The resulting bioluminescent dynamics upon chemical exposure have been demonstrated to reflect the treatment effects on cellular growth and metabolism, making these cells an inexpensive, continuous, real-time toxicity screening tool that can easily be adapted for high-throughput automation.

Mammalian  cell-based in vitro assays have been  widely employed as alternatives to animal testing for toxicological studies but  have been limited due to ...

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

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

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

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

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

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

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

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

Imaging G Protein-coupled Receptor-mediated Chemotaxis and its Signaling Events in Neutrophil-like HL60 Cells

Eukaryotic cells sense and move towards a chemoattractant gradient, a cellular process referred as chemotaxis. Chemotaxis plays critical roles in many physiological processes, such as embryogenesis, neuron patterning, metastasis of cancer cells, recruitment of neutrophils to sites of inflammation, and the development of the model organism Dictyostelium discoideum. Eukaryotic cells sense chemo-attractants using G protein-coupled receptors. Visual chemotaxis assays are essential for a better understanding of how eukaryotic cells control chemoattractant-mediated directional cell migration. Here, we describe detailed methods for: 1) real-time, high-resolution monitoring of multiple chemotaxis assays, and 2) simultaneously visualizing the chemoattractant gradient and the spatiotemporal dynamics of signaling events in neutrophil-like HL60 cells.

Visual chemotaxis assays are essential for a better understanding of how eukaryotic cells control chemoattractant-mediated directional cell migration. Here, we describe detailed methods for: 1) real-time, high-resolution monitoring of multiple chemotaxis assays, and 2) simultaneously visualizing the chemoattractant gradient and the spatiotemporal dynamics of signaling events in neutrophil-like HL60 cells.

Eukaryotic cells sense and move towards a chemoattractant gradient, a cellular process referred as chemotaxis. Chemotaxis plays critical roles in many ...

Identification of Intracellular Signaling Events Induced in Viable Cells by Interaction with Neighboring Cells Undergoing Apoptotic Cell Death

Cells dying by apoptosis, also referred to as regulated cell death, acquire multiple new activities that enable them to influence the function of adjacent live cells. Vital activities, such as survival, proliferation, growth, and differentiation, are among the many cellular functions modulated by apoptotic cells. The ability to recognize and respond to apoptotic cells appears to be a universal feature of all cells, regardless of lineage or organ of origination. However, the diversity and complexity of the response to apoptotic cells mandates that great care be taken in dissecting the signaling events and pathways responsible for any particular outcome. In particular, one must distinguish among the multiple mechanisms by which apoptotic cells can influence intracellular signaling pathways within viable responder cells, including: receptor-mediated recognition of the apoptotic cell, release of soluble mediators by the apoptotic cell, and/or engagement of the phagocytic machinery. Here, we provide a protocol for identifying intracellular signaling events that are induced in viable responder cells following their exposure to apoptotic cells. A major advantage of the protocol lies in the attention it pays to dissection of the mechanism by which apoptotic cells modulate signaling events within responding cells. While the protocol is specific for a conditionally immortalized mouse kidney proximal tubular cell line (BU.MPT cells), it is easily adapted to cell lines that are non-epithelial in origin and/or derived from organs other than the kidney. The use of dead cells as a stimulus introduces several unique factors that can hinder the detection of intracellular signaling events. These problems, as well as strategies to minimize or circumvent them, are discussed within the protocol. Application of this protocol should aid our expanding knowledge of the broad influence that dead or dying cells exert on their live neighbors, both in health and in disease.

Here, we present a protocol for the determination of intracellular signaling events induced in viable cells by physical interaction with adjacent dead or dying cells. The protocol focuses on signaling events induced by receptor-mediated recognition of the dead cells, as opposed to their phagocytic uptake or release of soluble mediators.

Cells dying by apoptosis, also referred to as regulated cell death, acquire multiple new activities that enable them to influence the function of adjacent ...

Characterization of Calcification Events Using Live Optical and Electron Microscopy Techniques in a Marine Tubeworm

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH distribution and calcium ions can be observed using live microscopy over time. This allows identification of the life stage and the tissue with the feature of interest for further electron microscopy studies. Life stage and tissues of interest are typically higher in pH and Ca signals. 
Here, using H. elegans, we present a protocol to characterize the presence of calcium carbonate structures in a biological specimen on the scanning electron microscope (SEM), using energy-dispersive X-ray spectroscopy (EDS) to visualize elemental composition, using electron backscatter diffraction (EBSD) to determine the presence of crystalline structures, and using transmission electron microscopy (TEM) to analyze the composition and structure of the material. In this protocol, a focused ion beam (FIB) is used to isolate samples with dimension suitable for TEM analysis. As FIB is a site specific technique, we demonstrate how information from the previous techniques can be used to identify the region of interest, where Ca signals are highest.

We demonstrate the use of various microscopy methods that are useful in observing the calcification of a tubeworm, Hydroides elegans, as well as locating and characterizing the first calcified material. Live microscopy and electron microscopy are used together to provide functional and material information that are important in studying biomineralization.

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH ...

Photostimulation by Femtosecond Laser Activates Extracellular-signal-regulated Kinase (ERK) Signaling or Mitochondrial Events in Target Cells

Direct control of cellular defined molecular events is important to life science. Recently, studies have demonstrated that femtosecond laser stimulation can simultaneously activate multiple cellular molecular signaling pathways. In this protocol, we show that through coupling femtosecond laser into a confocal microscope, cells can be stimulated precisely by the tightly-focused laser. Some molecular processes that can be simultaneously observed are subsequently activated. We present detailed protocols of the photostimulation to activate extracellular signal regulated kinase (ERK) signaling pathway in Hela cells. Mitochondrial flashes of reactive oxygen species (ROS) and other mitochondrial events can be also stimulated if focusing the femtosecond laser pulse on a certain mitochondrial tubular structure. This protocol includes pretreating cells before photostimulation, delivering the photostimulation by a femtosecond laser flash onto the target, and observing/identifying molecular changes afterwards. This protocol represents an all-optical tool for related biological researches.

Photostimulation by Femtosecond Laser Activates Extracellular-signal-regulated Kinase ERK Signaling or Mitochondrial Events in Target Cells

Tightly-focused femtosecond laser can deliver precise stimulation to cells by being coupled into a confocal microscopy enabling the real-time observation and photostimulation. The photostimulation can activate cell molecular events including ERK signaling pathway and mitochondrial flashes of reactive oxygen species.

Direct control of cellular defined molecular events is important to life science. Recently, studies have demonstrated that femtosecond laser stimulation ...