Name: measurement of force-sensitive protein dynamics in living cells using a combination of fluorescent techniques
Uploaded: 2018-11-02T15:00:00
Description: Watch this Scientific Journal Video about Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques at JoVE.com

This work was supported by a National Science Foundation CAREER Award (NSF-CMMI-14-54257) as well as grants from the American Heart Association (16GRNT30930019) and National Institutes of Health (R01GM121739-01) awarded to Dr. Brenton Hoffman and a National Science Foundation Graduate Research Fellowship awarded to Katheryn Rothenberg.&#160;The content is solely the responsibility of the authors and does not necessarily represent the official views of the NSF or NIH.

1. Generate Samples for Imaging
<ol>
	<li>Stably express tension sensor construct in desired cell type.
	<ol>
		<li>Clone tension sensor construct into pRRL vector or other viral expression plasmid. 
		NOTE: Several different molecular cloning tools are available to achieve this step including the use of restriction enzymes, overlap extension, and Gibson Assembly35. The pRRL vector is used in lenti viral transduction and enables a substantial degree of protein production through the use of ...

The FRET-FRAP method allows for direct measurement of force-sensitive protein dynamics, a property that has been difficult to directly probe inside living cells. The sensitivity of protein dynamics to molecular load is critical to the protein&#39;s function as a force transmitter or transducer. Loading is required for the transmission of both internally-generated and externally-applied forces, called mechanotransmission, and for the conversion of those forces into biochemically-detectable signals, called mechanotransduct...

The extracellular environment is a rich source of both biochemical and physical cues that dictate cell behavior. In particular, the physical nature of the microenvironment can mediate key cellular functions, including cell growth, migration, and differentiation1,2,3,4. Dysregulation of the mechanics of the microenvironment is a critical component to many diseases that do not yet have adequate treatments, such as cancer5, atherosclerosis6, and fibrosis7. A complete understanding of how cells convert physical stimuli into biochemically-detectable signals, a process termed mechanotransduction, requires the elucidation of the molecular mechanisms mediating force transmission, both into and out of the cells and within multiple subcellular structures.
Inside subcellular structures, proteins are constantly turning over; binding and unbinding based on the strength of their interactions with binding partners8. For forces to be successfully transmitted across a physical distance, there must be an unbroken chain of protein-protein interactions, meaning that a protein&#39;s turnover must be slow enough to sustain and transmit force to its binding partner9. While protein-protein interactions generally consist of several non-covalent bonds between the protein domains, the interaction is often conceptualized as a bound state that can transition to an unbound state under different conditions10,11. For a given protein-protein interaction, it is possible that force can have no effect on the lifetime of the interaction, known as an &#34;ideal bond&#34;, reduce the lifetime of the interaction, known as a &#34;slip bond&#34;, or increase the lifetime of the interaction, known as a &#34;catch bond&#34;10. Thus, there is an intricate relationship between protein load and protein dynamics, which we refer to as force-sensitive dynamics.
Towards understanding the effect of load on bond dynamics, a number of highly informative experiments have been performed on the single-molecule level. Using isolated proteins, or fragments of proteins and manipulation techniques such as magnetic tweezers, optical tweezers, and atomic force microscopy, these studies have demonstrated force-sensitive protein-protein interactions for several relevant proteins11,12. Both integrins13 and cadherins14, which are transmembrane proteins important for forming cell-matrix and cell-cell interactions, respectively, have demonstrated alterations in dynamics due to load. Within the cell, vinculin is recruited to both talin15 and &#945;-catenin16 in a force-dependent manner and can form a catch bond with actin17, indicating a crucial role for vinculin at both focal adhesions (FAs) and adherens junctions (AJs) under load. Single-molecule studies allow for the isolation of specific protein-protein interactions and yield unambiguous results, but they do not account for the complexity of the cellular environment.
Landmark experiments demonstrated that several subcellular structures, including FAs and AJs, are mechanosensitive, and exhibit enhanced assembly in response to internally-generated or externally-applied loads18,19,20,21,22. Additionally, several theoretical models have suggested that mechanosensitive assembly could be driven by force-sensitive protein dynamics23,24,25. To examine these force-sensitive dynamics within living cells, a few indirect approaches have been taken. FRAP and related techniques provide a relatively simple methodology for measuring protein dynamics in cells26,27,28,29. However, the measurement of protein load has been more limited. A typical approach is to compare protein dynamics in cells with and without the exposure to a cytoskeletal inhibitor used to reduce overall cell contractility8,30,31. Conceptually, this is a comparison between a high load and low load state. However, there is no quantification of the load across the protein in either state, and there may be unintended biochemical effects of the inhibitor, such the loss of key binding sites along an F-actin filament. Another approach, specific to FAs, has been to measure total force exertion on the substrate by the FA using traction force microscopy to approximate molecular load and examine the relationship with the dynamics of a single protein within the FA32. While this approach allows for the quantification of total force, it does not provide molecularly specific information. FAs are made up of over 200 different proteins, many of which can bear load33. Thus, measuring the total force output of an FA potentially obscures the possibility of multiple force transmission pathways and does not reliably provide a measure of load on a specific protein.
Unlike previous approaches in mechanobiology, the advent of FRET-based tension sensors allows direct measurement of loads experienced by specific proteins inside living cells34,35,36. Here, we present a protocol that combines FRET-based tension sensors with FRAP-based measure of protein dynamics. We refer to this technique as FRET-FRAP. This approach enables the simultaneous measurement of protein load and protein dynamics, thus allowing the assessment of the force-sensitive protein dynamics in living cells (Figure 1). Already, the FRET-FRAP technique has been applied to the study of the force-sensitive dynamics of the mechanical linker protein vinculin37. Tension sensors have been developed for numerous proteins that are relevant in a variety of subcellular structures. For example, sensors have been developed for vinculin34 and talin38,39 in FAs, cadherins and catenins in AJs40,41,42, nesprin in the nuclear LINC complex43, &#945;-actinin44 and filamin36 in the cytoskeleton, and MUC-1 in the glycocalyx45, among others46. Similarly, FRAP is a commonly used technique has been used on mechanosensitive proteins within the focal adhesions8,31, adherens junctions47, actin cortex26, and nucleus48. Moving forward, the FRET-FRAP technique should be broadly applicable to any of these existing sensors or newly developed sensors, allowing for the measurements of force-sensitive dynamics in a wide variety of subcellular structures and contexts. Towards this end, we provide a detailed, generalized protocol for implementing the FRET-FRAP technique applicable in these different systems. Hopefully, this will enable a wide variety of experiments elucidating the roles of various mechanosensitive proteins in regulating force transmission and in mediating cell behavior.

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			<td>0.05% Trypsin-EDTA</td>
			<td>Thermo Fisher</td>
			<td>25300062</td>
			<td></td>
		</tr>
		<tr>
			<td>16% Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>30525-89-4</td>
			<td></td>
		</tr>
		<tr>
			<td>60x Objective NA1.35</td>
			<td>Olympus</td>
			<td>UPLSAPO 60XO</td>
			<td></td>
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			<td>Antibiotic-Antimycotic Solution (100x)</td>
			<td>Gibco</td>
			<td>15240-062</td>
			<td></td>
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			<td>Automated Stage</td>
			<td>Prior Scientific</td>
			<td>H117EIX3</td>
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			<td>Custom Dichroic Mirror</td>
			<td>Chroma Technology Corp</td>
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			<td>T450/514rpc</td>
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			<td>Custom mTFP1 Emission Filter</td>
			<td>Chroma Technology Corp</td>
			<td></td>
			<td>ET485/20m</td>
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			<td>Custom mTFP1 Excitation Filter</td>
			<td>Chroma Technology Corp</td>
			<td></td>
			<td>ET450/30x</td>
		</tr>
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			<td>Custom Venus Excitation Filter</td>
			<td>Chroma Technology Corp</td>
			<td></td>
			<td>ET514/10x</td>
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			<td>DMEM-gfp Live Cell Visualization Medium</td>
			<td>Sapphire</td>
			<td>MC102</td>
			<td></td>
		</tr>
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			<td>Dulbecco&#39;s Modified Eagle&#39;s Medium&#160;</td>
			<td>Sigma Aldrich</td>
			<td>D5796</td>
			<td>with L-glutamine and sodium bicarbonate</td>
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			<td>Fetal Bovine Serum</td>
			<td>HyClone</td>
			<td>SH30396.03</td>
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			<td>Fibronectin, Human</td>
			<td>Corning</td>
			<td>47743-654</td>
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			<td>FRAPPA Calibration Slide</td>
			<td>Andor</td>
			<td></td>
			<td>provided along with FRAPPA unit</td>
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			<td>FRAPPA System with 515 nm Laser</td>
			<td>Andor</td>
			<td></td>
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			<td>Glass-bottomed Fluoro Dishes</td>
			<td>World Precision Instruments</td>
			<td>FD35</td>
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			<td>HEK293-T Cells</td>
			<td>ATCC</td>
			<td>CRL-3216</td>
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			<td>Hexadimethrine Bromide, Polybrene</td>
			<td>Sigma Aldrich</td>
			<td>H9268-5G</td>
			<td></td>
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			<td>High-glucose Dulbecco&#39;s Modified Eagle&#39;s Medium</td>
			<td>Sigma Aldrich</td>
			<td>D6429</td>
			<td></td>
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			<td>Inverted Fluorescent Microscope</td>
			<td>Olympus</td>
			<td>IX83</td>
			<td></td>
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		<tr>
			<td>JMP Pro Software</td>
			<td>SAS</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Lambda 10-3 Motorized Filter Wheels</td>
			<td>Sutter Instruments</td>
			<td>LB10-NW</td>
			<td></td>
		</tr>
		<tr>
			<td>LambdaLS Arc Lamp with 300W Ozone-Free Xenon Bulb</td>
			<td>Sutter Instruments</td>
			<td>LS/OF30</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine 2000</td>
			<td>Invitrogen</td>
			<td>11668-027</td>
			<td></td>
		</tr>
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			<td>MATLAB Software</td>
			<td>Mathworks</td>
			<td></td>
			<td></td>
		</tr>
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			<td>MEM Non-Essential Amino Acids</td>
			<td>Thermo Fisher</td>
			<td>11140050</td>
			<td></td>
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			<td>MetaMorph for Olympus</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
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			<td>Micro-Humidification System</td>
			<td>Bioptechs</td>
			<td>130708</td>
			<td></td>
		</tr>
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			<td>MoFlo Astrios EQ Cell Sorter</td>
			<td>Beckman Coulter</td>
			<td>B25982</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective Heater Medium</td>
			<td>Bioptechs</td>
			<td>150819-13</td>
			<td></td>
		</tr>
		<tr>
			<td>OptiMEM</td>
			<td>Thermo Fisher</td>
			<td>31985070</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline</td>
			<td>Sigma Aldrich</td>
			<td>D8537</td>
			<td></td>
		</tr>
		<tr>
			<td>pMD2.G Envelope Plasmid</td>
			<td>Addgene</td>
			<td>12259</td>
			<td></td>
		</tr>
		<tr>
			<td>pRRL Vector</td>
			<td></td>
			<td></td>
			<td>gift from Dr. Kam Leong (Columbia University)</td>
		</tr>
		<tr>
			<td>psPax2 Packaging Plasmid</td>
			<td>Addgene</td>
			<td>12260</td>
			<td></td>
		</tr>
		<tr>
			<td>sCMOS ORCA-Flash4.0 V2 Camera</td>
			<td>Hamamatsu Photonics</td>
			<td>C11440-22CU</td>
			<td></td>
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			<td>Sorvall Legend XT/XF Centrifuge</td>
			<td>Thermo Fisher</td>
			<td>75004505</td>
			<td></td>
		</tr>
		<tr>
			<td>Stable Z Stage Warmer</td>
			<td>Bioptechs</td>
			<td>403-1926</td>
			<td></td>
		</tr>
		<tr>
			<td>Venus Emission Filter</td>
			<td>Semrock</td>
			<td>FF01-571/72</td>
			<td></td>
		</tr>
	</tbody>
</table>

measurement of force-sensitive protein dynamics in living cells using a combination of fluorescent techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

FRET-FRAP is made up of the combination of two fluorescent techniques, FRET and FRAP. As we focused on the effects of protein load, we used FRET-based tension sensors34,46. These sensors are often based on a tension sensing module consisting of two fluorescent proteins, such as mTFP1 and VenusA206K, connected by a flagelliform linker (Figure 1A). When the module is placed between the head and tail dom...

Watch this Scientific Journal Video about Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques at JoVE.com

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

This method can answer key questions in the field of mechanobiology, such as how forces are transmitted and sensed within cells as well as between cells and their environment. The main advantage of this technique is it allows access to the molecular scale information regarding how proteins respond to mechanical forces inside the native context of the living cell. Though this method has been applied specifically to studying the focal adhesion protein vinculin, it can also be applied to other mechanically-sensitive proteins, such as talin, cadherins or nesprin.Visual demonstration of this method is critical, as proper imaging setup is difficult but necessary for the success of the analysis. Begin this procedure with stable expression of the tension sensor construct in the desired cell type, as detailed in the text protocol. To prepare substrates for cell seeding, acquire four 35 millimeter glass-bottomed dishes.Working in a cell culture hood, in a 15 milliliter conical tube, make four milliliters of 10 micrograms per milliliter fibronectin in sterile phosphate-buffered saline solution, or PBS. Gently invert the tube once to mix and let the solution sit for five minutes in the cell culture hood. Then pipette one milliliter of fibronectin solution onto each glass-bottomed dish.Leave the fibronectin solution on the dishes for one hour at room temperature or overnight at four degrees Celsius. Then aspirate the fibronectin solution and rinse once with PBS. Leave one milliliter of PBS on the dishes until the cells are ready for seeding.After counting the cells, seed 30, 000 cells onto each fibronectin-coated glass dish with the appropriate complete media for a final volume of 1.5 milliliters. Allow the cells to spread for four hours following seeding. At two hours of spreading, aspirate the growth media and rinse once with imaging media.Leave 1.5 milliliters of the imaging media. Warm up the microscope as described in the text protocol before beginning calibration. To calibrate the FRAP laser, first open the laser configuration window.Set Illumination setting to the appropriate FRAP Illumination settings for laser exposure to the sample. Then set Illumination setting to the illumination settings appropriate for imaging only the acceptor fluorophore. Select the objective to calibrate under Coordinate system setting.Uncheck Manually click calibration points and check Display images during calibration. Set the Dwell time to 10, 000 microseconds and the Number of pulses to 100. Now place the calibration slide, made of ethidium bromide sealed between a glass slide and a cover slip, into the stage adapter with the cover slip side down.Use the acceptor illumination settings to focus on the surface of the slide, identifiable as the focal plane with the brightest signal. Small defects in the coating will be visible to aid in focusing. Move the slide to an area with uniform fluorescence across the imaging plane.Click on Create Setting for the first calibration or Update Setting for subsequent calibrations. The software will initialize the calibration process, automatically bleaching and detecting the position of the bleached point. Ensure a successful calibration by assessing the final image, which will be a three-by-three grid of bleached points that should be evenly distributed and in focus.Save the calibration image for future reference. Remove the calibration slide and safely store. Calibration should be performed before beginning each experiment but does not need to be performed between samples.Prepare the microscopy setup for live cell imaging, preferably with a heated stage and objective, as well a carbon dioxide control. Allow to equilibrate for 20 minutes. Now place one of the generated samples of cells expressing the tension sensor into the microscope holder for imaging.Allow the cells to equilibrate for 10 minutes. To ensure the health of the imaged cells, maintain the temperature at 37 degrees Celsius in the imaging chamber. Use a peristaltic pump to pass humidified 5%carbon dioxide over the samples at 15 milliliters per minute to maintain the pH.Open the Multi-Dimensional Acquisition, or MDA, tool and set it up with FRET imaging parameters, including the different filter sets. Save this MDA to the Experimental Folder with the name of MDA_FRET_date. Set up another MDA with the FRAP imaging parameters, including the different filter sets, the Timelapse settings and the Journal to pulse the laser after pre-bleach acquisition.Save this MDA to the Experimental Folder with the name of MDA_FRAP_date. In the toolbar at the top of the screen, select Journal, Start Recording. Open the MDA window, load the MDA_FRET_date state and press Acquire.Then load the MDA_FRAP_date state and press Acquire. At the end of the acquisition, in the toolbar at the top of the screen, select Journal, Stop Recording. Save this Journal to the Experimental Folder with the name of FRETFRAP_date and add it to a toolbar for easy access.Close the MDA window. Navigate the sample using the image acquisition under Acquire, Acquire with minimal exposure time and neutral density filter to identify the cells of interest. Set continuous autofocus by navigating to Devices, Focus.Manually focus on the sample until reaching the correct imaging plane. Click Set Continuous Focus and wait for the system to adjust. Once the system adjusts, click Start Continuous Focusing.Then find a cell expressing the tension sensor with clear localization to a structure of interest and snap an image. Draw a rectangular region of interest, or ROI, to highlight where to bleach. Store this ROI location.Now initialize the FRETFRAP_date journal, which will begin the acquisition of FRET images, followed by the initialization of the FRAP timelapse. Repeat these steps until 10 to 15 image sets are acquired. Then analyze the FRET-FRAP data as detailed in the text protocol.Shown here are representative results for FRET imaging of the vinculin tension sensor, following segmentation and masking to isolate focal adhesions. Spacial variation of vinculin load can be seen across the cell. FRAP imaging and analysis can be affected by focal adhesion stability.A focal adhesion that is translocating rapidly during the imaging period can be difficult to accurately track. Often this is indicated by a FRAP curve that contains discontinuities and in uninterpretable. For a wild-type vinculin, there is an inverse correlation between protein load and turnover rate, indicating that vinculin is stabilized by force.Introducing a mutation in vinculin to effect its binding to talin results in a reversal of the correlation, indicating that vinculin that is unable to bind talin is destabilized by force. While attempting this procedure, it's important to remember to prepare samples properly, ensure that cells are expressing the sensor at endogenous levels and choose imaging parameters carefully. Following this procedure, other methods like immunofluorescence, measuring cellular scale dynamics or challenging cells with various inhibitors can be performed in order to answer additional questions like how the protein of interest interacts with other subcellular components to coordinate response to force.

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Fluorescent Nanoparticles for the Measurement of Ion Concentration in Biological Systems

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion fluxes at the cellular level are required for initiating action potentials in excitable cells.2 Sodium regulation plays an important role in both of these cases; however, no method currently exists for continuously monitoring sodium levels in vivo 3 and intracellular sodium probes 4 do not provide similar detailed results as calcium probes. In an effort to fill both of these voids, fluorescent nanosensors have been developed that can monitor sodium concentrations in vitro and in vivo.5,6 These sensors are based on ion-selective optode technology and consist of plasticized polymeric particles in which sodium specific recognition elements, pH-sensitive fluorophores, and additives are embedded.7-9 Mechanistically, the sodium recognition element extracts sodium into the sensor. 10 This extraction causes the pH-sensitive fluorophore to release a hydrogen ion to maintain charge neutrality within the sensor which causes a change in fluorescence. The sodium sensors are reversible and selective for sodium over potassium even at high intracellular concentrations.6 They are approximately 120 nm in diameter and are coated with polyethylene glycol to impart biocompatibility. Using microinjection techniques, the sensors can be delivered into the cytoplasm of cells where they have been shown to monitor the temporal and spatial sodium dynamics of beating cardiac myocytes.11 Additionally, they have also tracked real-time changes in sodium concentrations in vivo when injected subcutaneously into mice.3 Herein, we explain in detail and demonstrate the methodology for fabricating fluorescent sodium nanosensors and briefly demonstrate the biological applications our lab uses the nanosensors for: the microinjection of the sensors into cells; and the subcutaneous injection of the sensors into mice.

Fluorescent nanoparticles produced in our lab are used for imaging ion concentrations and ion fluxes in biological systems such as cells during signaling and interstitial fluid during physiological homeostasis.

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion ...

Fluorescence Lifetime Imaging of Molecular Rotors in Living Cells

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular events. Viscosity is one of the key parameters affecting the diffusion of molecules and proteins, and changes in viscosity have been linked to disease and malfunction at the cellular level.1-3 While methods to measure the bulk viscosity are well developed, imaging microviscosity remains a challenge. Viscosity maps of microscopic objects, such as single cells, have until recently been hard to obtain. Mapping viscosity with fluorescence techniques is advantageous because, similar to other optical techniques, it is minimally invasive, non-destructive and can be applied to living cells and tissues.
Fluorescent molecular rotors exhibit fluorescence lifetimes and quantum yields which are a function of the viscosity of their microenvironment.4,5 Intramolecular twisting or rotation leads to non-radiative decay from the excited state back to the ground state. A viscous environment slows this rotation or twisting, restricting access to this non-radiative decay pathway. This leads to an increase in the fluorescence quantum yield and the fluorescence lifetime. Fluorescence Lifetime Imaging (FLIM) of modified hydrophobic BODIPY dyes that act as fluorescent molecular rotors show that the fluorescence lifetime of these probes is a function of the microviscosity of their environment.6-8 A logarithmic plot of the fluorescence lifetime versus the solvent viscosity yields a straight line that obeys the F&ouml;rster Hoffman equation.9 This plot also serves as a calibration graph to convert fluorescence lifetime into viscosity.
Following incubation of living cells with the modified BODIPY fluorescent molecular rotor, a punctate dye distribution is observed in the fluorescence images. The viscosity value obtained in the puncta in live cells is around 100 times higher than that of water and of cellular cytoplasm.6,7 Time-resolved fluorescence anisotropy measurements yield rotational correlation times in agreement with these large microviscosity values. Mapping the fluorescence lifetime is independent of the fluorescence intensity, and thus allows the separation of probe concentration and viscosity effects. 
In summary, we have developed a practical and versatile approach to map the microviscosity in cells based on FLIM of fluorescent molecular rotors.

Fluorescence Lifetime Imaging (FLIM) has emerged as a key technique to image the environment and interaction of specific proteins and dyes in living cells. FLIM of fluorescent molecular rotors allows mapping of viscosity in living cells.

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular ...

Creating Two-Dimensional Patterned Substrates for Protein and Cell Confinement

Microcontact printing provides a rapid, highly reproducible method for the creation of well-defined patterned substrates.1 While microcontact printing can be employed to directly print a large number of molecules, including proteins,2 DNA,3 and silanes,4 the formation of self-assembled monolayers (SAMs) from long chain alkane thiols on gold provides a simple way to confine proteins and cells to specific patterns containing adhesive and resistant regions. This confinement can be used to control cell morphology and is useful for examining a variety of questions in protein and cell biology. Here, we describe a general method for the creation of well-defined protein patterns for cellular studies.5 This process involves three steps: the production of a patterned master using photolithography, the creation of a PDMS stamp, and microcontact printing of a gold-coated substrate. Once patterned, these cell culture substrates are capable of confining proteins and/or cells (primary cells or cell lines) to the pattern.
The use of self-assembled monolayer chemistry allows for precise control over the patterned protein/cell adhesive regions and non-adhesive regions; this cannot be achieved using direct protein stamping. Hexadecanethiol, the long chain alkane thiol used in the microcontact printing step, produces a hydrophobic surface that readily adsorbs protein from solution. The glycol-terminated thiol, used for backfilling the non-printed regions of the substrate, creates a monolayer that is resistant to protein adsorption and therefore cell growth.6 These thiol monomers produce highly structured monolayers that precisely define regions of the substrate that can support protein adsorption and cell growth. As a result, these substrates are useful for a wide variety of applications from the study of intercellular behavior7 to the creation of microelectronics.8
While other types of monolayer chemistry have been used for cell culture studies, including work from our group using trichlorosilanes to create patterns directly on glass substrates,9 patterned monolayers formed from alkane thiols on gold are straight-forward to prepare. Moreover, the monomers used for monolayer preparation are commercially available, stable, and do not require storage or handling under inert atmosphere. Patterned substrates prepared from alkane thiols can also be recycled and reused several times, maintaining cell confinement.10

Self-assembled monolayers (SAMs) formed from long chain alkane thiols on gold provide well-defined substrates for the formation of protein patterns and cell confinement. Microcontact printing of hexadecanethiol using a polydimethylsiloxane (PDMS) stamp followed by backfilling with a glycol-terminated alkane thiol monomer produces a pattern where protein and cells adsorb only to the stamped hexadecanethiol region.

Microcontact printing provides a rapid, highly reproducible method for the creation of well-defined patterned substrates.1  While microcontact printing ...

Spatio-Temporal Manipulation of Small GTPase Activity at Subcellular Level and on Timescale of Seconds in Living Cells

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular functions and events1, 2. Their spatiotemporally dynamic nature has been revealed by visualization of their activity and localization in real time3. In order to gain deeper understanding of their roles in diverse cellular functions at the molecular level, the next step should be perturbation of protein activities at a precise subcellular location and timing. 
To achieve this goal, we have developed a method for light-induced, spatio-temporally controlled activation of small GTPases by combining two techniques: (1) rapamycin-induced FKBP-FRB heterodimerization and (2) a photo-caging method of rapamycin. With the use of rapamycin-mediated FKBP-FRB heterodimerization, we have developed a method for rapidly inducible activation or inactivation of small GTPases including Rac4, Cdc424, RhoA4 and Ras5, in which rapamycin induces translocation of FKBP-fused GTPases, or their activators, to the plasma membrane where FRB is anchored. For coupling with this heterodimerization system, we have also developed a photo-caging system of rapamycin analogs. A photo-caged compound is a small molecule whose activity is suppressed with a photocleavable protecting group known as a caging group. To suppress heterodimerization activity completely, we designed a caged rapamycin that is tethered to a macromolecule such that the resulting large complex cannot cross the plasma membrane, leading to virtually no background activity as a chemical dimerizer inside cells6. Figure 1 illustrates a scheme of our system. With the combination of these two systems, we locally recruited a Rac activator to the plasma membrane on a timescale of seconds and achieved light-induced Rac activation at the subcellular level6.

A method for spatio-temporal control of small GTPase activity by light is described. This method is based on rapamycin-induced FKBP-FRB heterodimerization and photo-caging systems. Optimization of light-irradiation enables the spatio-temporally controlled activation of small GTPases at the subcellular level.

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular ...

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Ratiometric Biosensors that Measure Mitochondrial Redox State and ATP in Living Yeast Cells

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell death. These processes affect and are affected by the redox status of and ATP production by mitochondria. Here, we describe the use of two ratiometric, genetically encoded biosensors that can detect mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells. Mitochondrial redox state is measured using redox-sensitive Green Fluorescent Protein (roGFP) that is targeted to the mitochondrial matrix. Mito-roGFP contains cysteines at positions 147 and 204 of GFP, which undergo reversible and environment-dependent oxidation and reduction, which in turn alter the excitation spectrum of the protein. MitGO-ATeam is a F&ouml;rster resonance energy transfer (FRET) probe in which the &epsilon; subunit of the FoF1-ATP synthase is sandwiched between FRET donor and acceptor fluorescent proteins. Binding of ATP to the &epsilon; subunit results in conformation changes in the protein that bring the FRET donor and acceptor in close proximity and allow for fluorescence resonance energy transfer from the donor to acceptor.

We describe the use of two ratiometric, genetically encoded biosensors, which are based on GFP, to monitor mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells.

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell ...

Optical Control of Living Cells Electrical Activity by Conjugated Polymers

Hybrid interfaces between organic semiconductors and living tissues represent a new tool for in-vitro and in-vivo applications. In particular, conjugated polymers display several optimal properties as substrates for biological systems, such as good biocompatibility, excellent mechanical properties, cheap and easy processing technology, and possibility of deposition on light, thin and flexible substrates. These materials have been employed for cellular interfaces like neural probes, transistors for excitation and recording of neural activity, biosensors and actuators for drug release. Recent experiments have also demonstrated the possibility to use conjugated polymers for all-optical modulation of the electrical activity of cells. Several in-vitro study cases have been reported, including primary neuronal networks, astrocytes and secondary line cells. Moreover, signal photo-transduction mediated by organic polymers has been shown to restore light sensitivity in degenerated retinas, suggesting that these devices may be used for artificial retinal prosthesis in the future. All in all, light sensitive conjugated polymers represent a new approach for optical modulation of cellular activity.
In this work, all the steps required to fabricate a bio-polymer interface for optical excitation of living cells are described. The function of the active interface is to transduce the light stimulus into a modulation of the cell membrane potential. As a study case, useful for in-vitro studies, a polythiophene thin film is used as the functional, light absorbing layer, and Human Embryonic Kidney (HEK-293) cells are employed as the biological component of the interface. Practical examples of successful control of the cell membrane potential upon stimulation with light pulses of different duration are provided. In particular, it is shown that both depolarizing and hyperpolarizing effects on the cell membrane can be achieved depending on the duration of the light stimulus. The reported protocol is of general validity and can be straightforwardly extended to other biological preparations.

A method for stimulation of in-vitro cell cultures electrical activity with visible light, based on the use of organic semiconducting polymers is described.

Hybrid interfaces between organic semiconductors and living tissues represent a new tool for in-vitro and in-vivo applications. In particular, conjugated ...

Using Synthetic Biology to Engineer Living Cells That Interface with Programmable Materials

We have developed an abiotic-biotic interface that allows engineered cells to control the material properties of a functionalized surface. This system is made by creating two modules: a synthetically engineered strain of E. coli cells and a functionalized material interface. Within this paper, we detail a protocol for genetically engineering selected behaviors within a strain of E. coli using molecular cloning strategies. Once developed, this strain produces elevated levels of biotin when exposed to a chemical inducer. Additionally, we detail protocols for creating two different functionalized surfaces, each of which is able to respond to cell-synthesized biotin. Taken together, we present a methodology for creating a linked, abiotic-biotic system that allows engineered cells to control material composition and assembly on nonliving substrates.

This paper presents a series of protocols for developing engineered cells and functionalized surfaces that enable synthetically engineered E. coli to control and manipulate programmable material surfaces.

We have developed an abiotic-biotic interface that allows engineered cells to control the material properties of a functionalized surface. This system is ...

Production of Elastin-like Protein Hydrogels for Encapsulation and Immunostaining of Cells in 3D

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue culture systems lack a three-dimensional (3D) matrix, resulting in a significant disconnect between results collected in vitro and in vivo. To address this limitation, researchers have engineered 3D hydrogel tissue culture platforms that can mimic the biochemical and biophysical properties of the in vivo cell microenvironment. This research has motivated the need to develop material platforms that support 3D cell encapsulation and downstream biochemical assays. Recombinant protein engineering offers a unique toolset for 3D hydrogel material design and development by allowing for the specific control of protein sequence and therefore, by extension, the potential mechanical and biochemical properties of the resultant matrix. Here, we present a protocol for the expression of recombinantly-derived elastin-like protein (ELP), which can be used to form hydrogels with independently tunable mechanical properties and cell-adhesive ligand concentration. We further present a methodology for cell encapsulation within ELP hydrogels and subsequent immunofluorescent staining of embedded cells for downstream analysis and quantification.

Recombinant protein-engineered hydrogels are advantageous for 3D cell culture as they allow for complete tunability of the polymer backbone and therefore, the cell microenvironment. Here, we describe the process of recombinant elastin-like protein purification and its application in 3D hydrogel cell encapsulation.

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue ...