This work was supported by the Thomas F. and Kate Miller Jeffress Memoria Trust (to DEC) and NIH grant R35GM119617 (to DEC). The confocal microscope imaging was performed at the VCU Nanomaterials Characterization Core (NCC) Facility.

1. Obtain Nesprin-2G Sensor DNA and Other Plasmid DNA
<ol>
	<li>Obtain Nesprin-2G TS (tension sensor), Nesprin-2G HL (headless) control, mTFP1, venus, and TSmod from a commercial source. Propagate all the DNA plasmids and purify them using standard E. coli strains, such as DH5-&#945;, as described previously12,13.</li>
</ol>
2. Transfect Cells with Nesprin-2G and Other Plasmid DNA
<ol>
	<li>Grow NIH 3T3 fibroblasts cells to 70-90% confluence in a 6-well cell culture dish in a standard cell culture incubator with temperature (37 &#176;C) and CO2 (5%) regulation. For the cell growth medium, use Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal calf serum.</li>
	<li>In a cell culture hood, remove the medium and rinse each well with approximately 1 mL of a reduced-serum cell medium. Add 800 &#181;L of reduced-serum cell medium to each well and place the 6-well chamber in the incubator.</li>
	<li>Pipette 700 &#181;L of reduced-serum cell medium into a 1.5 mL tube with 35 &#181;L of lipid carrier solution to form the &#34;lipomix&#34;. Mix by pipetting. Label the tube with an &#34;L.&#34;</li>
	<li>Gather six 1.5 mL tubes and label them 1 through 6. Pipette 100 &#181;L of reduced-serum cell medium into each tube.
	<ol>
		<li>Using the plasmid DNA concentration from step 1, pipette 2 &#181;g of Nesprin 2G-TS into tubes 1 and 2. Pipette 2 &#181;g of Nesprin-HL into tubes 3 and 4. Pipette 1 &#181;g of mTFP into tube 5. Pipette 1 &#181;g of mVenus into tube 6. Do not re-use pipette tips when pipetting different types of DNA.</li>
	</ol>
	</li>
	<li>Pipette 100 &#181;L of the lipomix from the &#34;L&#34; tube into each labeled tube (1-6) and mix by repeated pipetting. Use a clean pipette for each tube. Incubate for 10-20 min.</li>
	<li>Add 200 &#181;L from each labeled tube to a well in the 6-well chamber with 70-90% confluent cells. Label the top of each well with the corresponding DNA added. Place the 6-well chamber in an incubator for 4-6 h.</li>
	<li>Aspirate the medium and add 1-2 mL of reduced-serum cell medium to rinse. Aspirate the reduced-serum cell medium, add 2 mL of trypsin to each well, and place the 6-well dish in the incubator (5-15 min).</li>
	<li>While the cells detach in the incubator, coat 6 glass-bottom viewing dishes with a layer of fibronectin at a concentration of 20 &#181;g/mL dissolved in phosphate-buffered saline (PBS). Allow the dishes to coat the surface in the cell culture hood (approximately 20 min).</li>
	<li>Neutralize the trypsin by adding 2 mL of DMEM once the cells are detached.</li>
	<li>Transfer the contents of each well to a labeled 15-mL centrifuge tube and spin down at 90 x g for 5 min in a swinging rotor centrifuge. Aspirate the supernatant and re-suspend each cell pellet in 1,000 &#181;L of DMEM by pipette mixing.</li>
	<li>Aspirate the fibronectin mixture from the glass dishes and pipette 1,000 &#181;L of each cell suspension onto the glass dishes.</li>
	<li>After the cells settle to the bottom of the glass dishes (~15 min), add another 1 mL of DMEM + 10% FBS + 1% Pen-Strep to each well and place in the cell incubator. Allow the cells to attach and express sensor for at least 18-24 h. 
	NOTE: Cells are transfected using commercial cationic lipid transfection reagents (see the Table of Materials). Alternatively, stable cell lines can be selected by using a plasmid with a gene conferring resistance to a toxin (the pcDNA plasmids for Nesprin-TS and -HL are available on a DNA repository website (see the Table of Materials) provide cells expressing geneticin resistance). Additionally, viral infection methods (lentivirus, retrovirus, or adenovirus) can be utilized to express the sensor in cells that are hard to transfect.</li>
	<li>In addition to Nesprin-TS, transfect additional cells with the Nesprin HL zero-force control, as described in steps 2.1-2.12; TSmod can also be used as a zero-force control and should exhibit similar FRET to Nesprin-HL.</li>
	<li>Transfect cells with mTFP1 and venus to generate spectral fingerprints (see step 4). 
	NOTE: mTFP1 and venus typically express at higher levels than Nesprin-TS, and as such, lower amounts of DNA may need to be transfected to achieve similar expression levels.</li>
</ol>
3. Verify Transfection Efficiency
<ol>
	<li>Roughly 18-24 h after completing the transfection, use an inverted fluorescent microscope to examine the efficiency of transfection by comparing the number of fluorescent cells to the total number of cells in view (usually 5-30%).
	<ol>
		<li>Use a fluorescent microscope equipped with an excitation frequency near 462 nm (mTFP1) or 525 nm (venus) and an emission filter centered near 492 nm (mTFP1) or 525 nm (venus). 
		NOTE: Alternatively, GFP filter sets will capture a combination of mTFP1 and venus emissions and will allow the confirmation of the transfection efficiency.</li>
	</ol>
	</li>
	<li>Image live cells within 48 h after transfection.
	<ol>
		<li>Alternatively, fix cells in 4% paraformaldehyde (in PBS with calcium and magnesium) for 5 min, store in PBS, and view after 48 h8. 
		NOTE: Beyond 48 h, the signal quality and strength decays. Fixing the cells preserves their state, including the FRET being expressed8; however, cells should only be imaged in PBS, as mounting medium may affect FRET14. Fixed cells can only be compared to other fixed cells, as there may be a change in expression. 
		Caution: Paraformaldehyde is toxic. Wear appropriate personal protective equipment (PPE).</li>
	</ol>
	</li>
</ol>
4. Capture Spectral Fingerprints of mTFP1 and Venus Fluorophores for Spectral Unmixing
<ol>
	<li>In a cell culture hood, replace the cell medium with imaging medium (HEPES-buffered) supplemented with 10% fetal bovine serum.</li>
	<li>Place the viewing dishes in a temperature-controlled (37 &#176;C) confocal microscope stage.</li>
	<li>Place the glass viewing dish with mTFP1-transfected cells over the oil objective at 60X magnification with a numerical aperture of 1.4. 
	NOTE: The oil is used on a laser scanning microscope on the 60X objective to closely resemble the refractive index of the glass substrate. The oil is placed on top of the objective lens and comes in direct contact with the glass coverslip.</li>
	<li>Locate mTFP1 expressing cells with a 458 nm excitation source and an emission bandpass filter centered at 500 nm.</li>
	<li>With a fluorescent cell in the field of view, select the spectral detection mode (&#34;Lambda Mode&#34; in the software used here) and capture the spectral image (Figure 3-1); include all frequencies beyond 458 nm using 10 nm increments (Figure 3-3). Select a bright fluorescent region (ROI) on the cell (20-pixel radius). 
	NOTE: The spectral shape of the mTFP1-expressing ROI should remain relatively constant across the cell. If the shape varies considerably, re-adjust the laser and gain settings to improve the signal-to-noise ratio.</li>
	<li>Add the fluorescent ROI mean, normalized intensity to the spectral database by clicking &#34;save spectra to database.&#34;</li>
	<li>Optimize the laser power and gain such that a good signal-to-noise ratio is achieved. Settings will vary for different equipment. Using non-fluorescent, untransfected cells as a background reference, increase the gain and power until the cells are bright, but not beyond the dynamic range of the detector (saturation point). Background cells should not have detectable fluorescence after averaging.</li>
	<li>Repeat the process for the venus-transfected cells with the following exceptions:
	<ol>
		<li>Ensure that the excitation frequency is at 515 nm and that the bandpass filter is centered near 530 nm when locating venus-expressing cells.</li>
		<li>In &#34;Spectral Mode,&#34; use an excitation frequency of 515 nm instead of 458 nm.</li>
	</ol>
	</li>
</ol>
5. Capture Unmixed Images
<ol>
	<li>Switch the capturing mode to &#34;Spectral Unmixing.&#34;</li>
	<li>Add the spectral fingerprints of venus and mTFP1 into the unmixing channels (Figure 3, Arrow-2).</li>
	<li>Set the excitation laser back to a 458 nm argon source.</li>
	<li>Place the Nesprin-TS viewing dish above the 60X oil objective.</li>
	<li>After focusing on a fluorescent cell with sufficiently bright expression, adjust the gain and laser power to optimize the signal-to-noise ratio. 
	NOTE: Since the FRET efficiency of Nesprin-TS is near 20%, the unmixed venus image should be substantially dimmer than the unmixed mTFP1 image. Once an acceptable power and gain setting have been determined iteratively, these parameters must remain constant for all images captured.</li>
	<li>Capture a minimum of 15-20 images of Nesprin-TS cells with relatively similar brightness and avoid excessive pixel saturation (all saturated pixels are removed during image processing).</li>
	<li>Repeat the image capturing process with Nesprin-HL cells using identical imaging parameters.</li>
</ol>
6. Image Processing and Ratio Image Analysis
<ol>
	<li>Using open-source ImageJ (FIJI) software (http://fiji.sc/), open native format images using BioFormats Reader.</li>
	<li>Pre-process the images and compute the ratio images using previously established protocols15.</li>
</ol>

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The authors have nothing to disclose.

A method and demonstration of live cell imaging of mechanical tension across Nesprin-2G, a protein in the nuclear LINC complex, was outlined above. Prior to this work, various techniques, such as micropipette aspiration, magnetic-bead cytometry, and microscopic laser-ablation, have been used to apply strain on the cell nucleus and to measure its bulk material properties16,17,18. However, until our recent work, no studies had directly shown that tensile forces are transferred directly onto an LINC complex protein in live cells2.
Previously, our laboratory demonstrated that a modified version of Nesprin-2G, known as mini-Nesprin2G, could be engineered to express a FRET-force probe known as TSmod (Figure 2C)2. TSmod was previously shown to dynamically measure tensile forces in live cells and in various load-bearing proteins3,4,6,8. It was further shown that mini-Nesprin-2G was indistinguishable from endogenous Nesprin-2G with respect to its localization, known effects on the cytoskeleton, and ability to rescue nuclear positioning11,19. Using the mini Nesprin-2G protein with the TSmod (Nesprin-TS), it was shown that NIH3T3 fibroblasts expressing the sensor have a baseline level of tension relative to Nesprin-HL, which cannot bind to actin2. Furthermore, the modulation of cellular contractility using inhibitors or activators of myosin can be detected by Nesprin-TS2.
There are a number of critical steps in this protocol. Confusing Nesprin-TS with Nesprin-HL (DNA or expressing cells) is not readily detected, as both similarly localize to the nuclear membrane, and will lead to confusing results. Since Nesprin-HL is a no-force control, the measured FRET should be higher than Nesprin-TS on average. To be cautious, sequencing isolated DNA constructs and carefully labeling cells is advised. Second, it is important to carefully optimize the detector gain and laser power on the confocal microscope to improve the signal-to-noise ratio and to minimize bleaching. To properly measure FRET, these parameters must remain constant during the image acquisition. Furthermore, optimizing the detector gain or power for dim or very bright expressing cells is suboptimal, because median expressing cells will tend to fall out of the dynamic range of the detector. Certain cells that express a high concentration of sensor tend to fluoresce in the nucleoplasm and endoplasmic reticulum, making it difficult to resolve the nuclear membrane, which is presumably the more interesting region with regards to force. Selecting cells with slightly lower expression levels tends to resolve this issue of sensor localization.
While ratiometric spectral imaging is a relatively simple and quick way to measure FRET, it does not output units of FRET efficiency due to acceptor signal bleed-through20. Without units of FRET efficiency, it is not possible to convert the FRET ratio index into a calibrated force measurement. Due to this limitation, only relative quantitative force comparisons can be made. FRET efficiency can be determined by more complicated methods, such as fluorescence-lifetime imaging microscopy (FLIM) or acceptor photobleaching. Absolute force (at the pN level) can be estimated from FRET efficiency, as previously described3,8.
In contrast to previous methods to measure the mechanical properties of the nucleus or the displacement of the nucleus based on externally applied forces, measuring the FRET from Nesprin-TS has the advantage of non-invasively probing tension on a component of the nuclear membrane16,17,18. The sensor outputs a fluorescent signal that is correlated with the strain on individual Nesprin-2G molecules. In contrast, micropipette aspiration requires the use of delicate tools when working with live cells. Confocal laser ablation causes local cellular damage and requires a pulsed laser source. Magnetic twisting cytometry transfers forces onto the nuclear membrane via the cytoskeleton and cannot be used to measure forces on specific proteins, unless it were to be coupled with the Nesprin-2G tension sensor.
While it has been demonstrated that actin-based tensile forces are transferred to Nesprin-2G, a component of the LINC complex, it remains unknown how much tensile or compressive forces are exerted onto other structural LINC proteins, such SUN-1 or SUN-2. Future work will be directed at cloning FRET-force probes into these putative load-bearing LINC proteins to further elucidate nuclear mechanics with protein-level resolution. Another issue that presents itself is that changes in FRET may be unrelated to changes in tension but rather reflect either changes in Nesprin-2G oligomerization or conformational changes of Nesprin-2G. Changes in cellular pH can influence the fluorescence of the sensor and change the measured FRET. The Nesprin-2G HL sensor represents a good control for some of these changes (as FRET changes observed with this sensor are most likely unrelated to force), and intermolecular FRET control constructs can be developed to assess oligomerization (see below). Additionally, the expression of nesprin-2G TS is driven by a CMV promoter, which results in overexpression, and this high level of expression may potentially induce biological artifacts. Recent advances with clustered regularly interspaced short palindromic repeats (CRISPR) have allowed for homology-directed repair (HDR) approaches to insert larger DNA sequences into the genome21. This approach could be used to modify endogenous Nesprin-2G (or other proteins) to include TSmod. This would provide expression of the force sensor using the appropriate endogenous promotor, reducing the potential for overexpression artifacts.
TSmod and other elastic FRET probes can also be used to develop new sensors for other proteins, which could then be used in a similar protocol as that described in this article. A major concern with the development of any new force sensor is determining the internal insertion site for the FRET sensor. First, the site of insertion must be in a region of the protein subjected to mechanical load. These regions can be identified by examining where cytoskeletal-connected proteins interact with the protein. Second, the insertion of the large (~50 kDa) TSmod must not disrupt the biological function of the protein being studied. A previously developed &#34;mini&#34; form of Nesprin-2G showed that actin-binding CH domains bridged to the SUN-binding KASH domain were sufficient for Nesprin-2G nuclear movement in wounded monolayers11,19, suggesting that the insertion of TSmod would likely not disrupt protein function. Confirmation of the biological function of the sensor requires a functional assay for protein function (in which the force sensor can be shown to rescue a specific function in cells depleted of endogenous protein). Third, changes in protein oligomerization can alter the FRET between proteins (termed intermolecular FRET22), which would result in FRET changes that are not related to force. A careful analysis of this often requires specialized intermolecular FRET constructs that specifically report oligomerization.
FRET-based force sensors have enabled the spatial and temporal analysis of mechanical forces with protein-specific resolution. Although this protocol details the use of the Nesprin-2G tension sensor, it is applicable to a number of existing sensors available for proteins in focal adhesions and cell-cell junctions. The use of these sensors for Nesprin-2G and other load-bearing proteins will provide significant insight into cell mechanics and mechanobiology.

Force-sensitive, genetically encoded FRET sensors have recently emerged as an important tool for measuring tensile-based forces in live cells, providing insight into how mechanical forces are applied across proteins1,2,3,4. With these tools, researchers can non-invasively image intracellular forces in living cells using conventional fluorescent microscopes. These sensors consist of a FRET-pair (donor and acceptor fluorescent proteins, most frequently a blue donor and yellow acceptor) separated by an elastic peptide3. In contrast to C- or N-terminal tagging, this sensor is inserted into an internal site of a protein to measure the mechanical force transmitted across the protein, behaving as a molecular strain gauge. Increased mechanical tension across the sensor results in an increased distance between the FRET-pair, resulting in decreased FRET3. As a result, the FRET is inversely related to tensile force.
These fluorescent-based sensors have been developed for focal adhesion proteins (vinculin3 and talin4), cytoskeletal proteins (&#945;-actinin5), and cell-cell junction proteins (E-Cadherin6,7, VE-Cadherin8, and PECAM8). The most frequently used and well-characterized elastic linker in these biosensors is known as TSmod and consists of a repetitive sequence of 40 amino acids, (GPGGA)8, which was derived from the spider silk protein flagelliform. TSmod has been shown to behave as a linear elastic nano-spring, with FRET responsiveness to 1 to 5 pN of tensile force3. Different lengths of flagelliform can be used to alter the dynamic range of TSmod FRET-force sensitivity9. In addition to flagelliform, spectrin repeats5 and villin headpiece peptide (known as HP35)4 have been used as the elastic peptides between FRET-pairs in similar force biosensors4. Finally, a recent report showed that TSmod can also be used to detect compressive forces10.
We recently developed a force sensor for the linker of the nucleo-cytoskeleton (LINC) complex protein Nesprin-2G by using TSmod inserted into a previously developed truncated Nesprin-2G protein known as mini-Nesprin2G (Figure 2C), which behaves similarly to endogenous Nesprin-2G11. The LINC complex contains multiple proteins that lead from the outside to the inside of the nucleus, linking the cytoplasmic cytoskeleton to the nuclear lamina. Nesprin-2G is a structural protein binding to both the actin cytoskeleton in the cytoplasm and to SUN proteins in the perinuclear space. Using our biosensor, we were able to show that Nesprin-2G is subject to actomyosin-dependent tension in NIH3T3 fibroblasts2. This was the first time that force was directly measured across a protein in the nuclear LINC complex, and it is likely to become an important tool to understand the role of force on the nucleus in mechanobiology.
The protocol below provides a detailed methodology of how to use the Nesprin-2G force sensor, including the expression of the Nesprin tension sensor (Nesprin-TS) in mammalian cells, as well as the acquisition and analysis of FRET images of cells expressing Nesprin-TS. Using an inverted confocal microscope equipped with a spectral detector, a description of how to measure sensitized emission FRET using spectral unmixing and ratiometric FRET imaging is provided. The output ratiometric images can be used to make relative quantitative force comparisons. While this protocol is focused on the expression of Nesprin-TS in fibroblasts, it is easily adaptable to other mammalian cells, including both cell lines and primary cells. Furthermore, this protocol as it relates to image acquisition and FRET analysis can readily be adapted to other FRET-based force biosensors that have been developed for other proteins.

<table><tbody><tr><td>Nesprin-TS DNA</td><td>Addgene</td><td>68127</td><td>Retrieve from https://www.addgene.org/68127/</td></tr><tr><td>Nesprin-HL DNA</td><td>Addgene</td><td>68128</td><td>Retrieve from https://www.addgene.org/68128/</td></tr><tr><td>mTFP1 DNA</td><td>Addgene</td><td>54613</td><td>Retrieve from https://www.addgene.org/54613/</td></tr><tr><td>mVenus DNA</td><td>Addgene</td><td>27793</td><td>Retrieve from https://www.addgene.org/27793/</td></tr><tr><td>TSmod DNA</td><td>Addgene</td><td>26021</td><td>Retrieve from https://www.addgene.org/26021/</td></tr><tr><td>Competent Cells</td><td>Bioline</td><td>BIO-85026</td><td></td></tr><tr><td>Liquid LB Media</td><td>ThermoFisher</td><td>10855001</td><td>https://www.thermofisher.com/order/catalog/product/10855001</td></tr><tr><td>Solid LB Bacterial Culture Plates</td><td>Sigma-Aldrich</td><td>L5667</td><td>http://www.sigmaaldrich.com/catalog/product/sigma/l5667?lang=en&amp;amp;region=US</td></tr><tr><td>Ampicillin</td><td>Sigma</td><td>A9518</td><td></td></tr><tr><td>Spectrophotometer</td><td>Biorad</td><td>273 BR 07335</td><td>SmartSpec Plus</td></tr><tr><td>quartz cuvette</td><td>Biorad</td><td>1702504</td><td>Cuvette for SmartSpec Plus</td></tr><tr><td>DNA isolation kit</td><td>Macherey-Nagel</td><td>740412.5</td><td>NucleoBond Xtra Midi Plus</td></tr><tr><td>6-well cell culture dish</td><td>Falcon-Corning</td><td>353046</td><td>Multiwell 6-well Polystrene Culture Dish</td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium, (DMEM) cell media</td><td>Gibco</td><td>11995-065</td><td>DMEM(1x)</td></tr><tr><td>Bovine Serum</td><td>Life Technologies</td><td>16170-078</td><td></td></tr><tr><td>reduced serum cell media</td><td>Gibco</td><td>31985-070</td><td>Reduced Serum Medium, &quot;optimem&quot;</td></tr><tr><td>Lipid Carrier Solution</td><td>invitrogen</td><td>11668-019</td><td>Lipid Reagent, &quot;Lipofectamine 2000&quot;</td></tr><tr><td>1.5 mL sterile plastic tube</td><td>Denville</td><td>c2170</td><td></td></tr><tr><td>Trypsin</td><td>Gibco</td><td>25200-056</td><td>0.25% Trypsin-EDTA (1x)</td></tr><tr><td>glass-bottom microscope viewing dish</td><td>In Vitro Scientific</td><td>D35-20-1.5-N</td><td>35 mm Dish with 20 mm Bottom Well #1.5 glass</td></tr><tr><td>Fibronectin</td><td>ThermoFisher</td><td>33016015</td><td>fibronectin human protein, plasma</td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>Gibco</td><td>14190-144</td><td>Dulbecco&#039;s Phosphate Buffered Saline</td></tr><tr><td>15 mL sterile centrifuge tube</td><td>Greiner bio-one</td><td>188261</td><td></td></tr><tr><td>swinging rotor centrifuge&amp;nbsp;</td><td>Thermo electron</td><td>Centra CL2</td><td>Swinging rotor thermo electron 236</td></tr><tr><td>cell culture biosafety hood</td><td>Forma Scientific</td><td>1284</td><td></td></tr><tr><td>climate controlled cell culture incubator</td><td>ThermoFisher</td><td>3596</td><td></td></tr><tr><td>inverted LED widefield fluorescent microscope</td><td>Life technologies</td><td>EVOS FL</td><td></td></tr><tr><td>Clear HEPES buffered imaging media</td><td>Molecular Probes</td><td>A14291DJ</td><td></td></tr><tr><td>Fetal bovine Serum</td><td>Life technologies</td><td>10437-028</td><td></td></tr><tr><td>Temperature Controlled-Inverted confocal w/458 and 515 nm laser sources&amp;nbsp;</td><td>Zeiss</td><td>&amp;nbsp;LSM 710-w/spectral META detector</td><td></td></tr><tr><td>Outgrowth Media</td><td>Newengland Biolabs</td><td>B9020s</td><td></td></tr><tr><td>NIH 3T3 Fibroblasts</td><td>ATCC</td><td>CRL-1658</td><td></td></tr></tbody></table>

a protocol for using f&#246;rster resonance energy transfer (fret)-force biosensors to measure mechanical forces across the nuclear linc complex

The LINC complex has been hypothesized to be the critical structure that mediates the transfer of mechanical forces from the cytoskeleton to the nucleus. Nesprin-2G is a key component of the LINC complex that connects the actin cytoskeleton to membrane proteins (SUN domain proteins) in the perinuclear space. These membrane proteins connect to lamins inside the nucleus. Recently, a F&#246;rster Resonance Energy Transfer (FRET)-force probe was cloned into mini-Nesprin-2G (Nesprin-TS (tension sensor)) and used to measure tension across Nesprin-2G in live NIH3T3 fibroblasts. This paper describes the process of using Nesprin-TS to measure LINC complex forces in NIH3T3 fibroblasts. To extract FRET information from Nesprin-TS, an outline of how to spectrally unmix raw spectral images into acceptor and donor fluorescent channels is also presented. Using open-source software (ImageJ), images are pre-processed and transformed into ratiometric images. Finally, FRET data of Nesprin-TS is presented, along with strategies for how to compare data across different experimental groups.

Following the protocol above, plasmid DNA was acquired from the DNA repository and transformed into E. coli cells. E. coli expressing the sensor DNA were selected from LB/Ampicillin plates and amplified in a liquid LB broth. Following the amplification of the vectors, DNA plasmids were purified into TRIS-EDTA buffer using a standard, commercially available DNA isolation kit. Using a spectrophotometer, purified DNA was quantified into a standard concentration of &#181;g/mL (Figure 1A, Days 1-2).
Using NIH3T3 fibroblasts, cells were grown to 70-90% confluence in a 6-well plastic cell culture dish. To insert plasmid DNA into the NIH3T3 cells, a lipid-mediated plasmid transfection was performed. Commercially available transfection reagent (see the Table of Materials) was used as the lipid vessel for DNA. Two replicates of the Nesprin-TS and Nesprin-HL sensors were transfected into 4 cell wells (Figure 1B). In preparation for FRET imaging, single fluorophore constructs of mTFP1 and venus were transfected into NIH3T3 cells (Figure 1B). Following transfection, cells were transferred to glass-bottom viewing dishes compatible with high magnification, inverted confocal microscopes.
Before proceeding to confocal imaging, a simple inverted wide-field fluorescent microscope was used to verify the transfection efficiency. Using a 10X objective with a numerical aperture of 0.25, many cells in a typical field-of-view expressed the Nesprin-TS (Figure 2). Generally, the sensor localized around the nuclear envelope, consistent with the endogenous localization of Nesprin-2G2. The no-force control sensor, Nesprin-HL, followed a similar expression pattern2.
After the confirmation of the transfection efficiency of Nesprin sensors TS and HL, live cells expressing single fluorophore constructs (mTFP1 or venus) were imaged with an inverted confocal microscope. Using the spectral detection mode, mTFP1 and venus spectral fingerprints were recorded in the spectral unmixing database (Figure 3, Arrow-2 and Figure 4A). Following fingerprinting, the capture parameter was switched to unmixing mode, with the mTFP1 and venus spectra loaded as the components (Figure 4D). Raw images were acquired as spectrally resolved image stacks (Figure 4).
Finally, the spectrally resolved image stacks were imported into ImageJ open-source software for processing. The images were pre-processed, and ratio images were computed using established procedures15.
Since the FRET ratio is a relative measurement, every experiment must compare two or more conditions. Additionally, because laser and camera detector settings can influence FRET, all conditions should be acquired in the same imaging session. Frequently, we use a force-insensitive headless or tailless control, which represents a zero-force control, as a base-line for our experiments. Because FRET ratio images can be noisy, it is often necessary to take multiple images (5-20) for each condition. For conditions where there are large differences in FRET, this will be visually discernable in the FRET ratio images. The most dramatic difference in FRET for the Nesprin tension sensor was between unpatterned and elongated cells (Figure 5). Although there is variation between cells, the change in FRET between the two conditions is so dramatic that further analysis may not be required. However, other conditions often have minimal changes in FRET; they may not be visually discernable between individual images, but rather require the data to be analyzed in aggregate. To determine if these changes are significant, the median FRET ratio for each image is computed and visually expressed as a set of histograms. By comparing histograms between conditions, it becomes easier to see subtler changes in FRET between experimental groups. In Figure 6A and 6B, the FRET histograms of each individual cell are represented by a color in the stacked histogram. Calyculin A-treated cells (Figure 6B) exhibit a leftward-shifted FRET histogram relative to cells treated with myosin inhibitor ML7 (Figure 6A).
<img alt="Figure 1" src="/files/ftp_upload/54902/54902fig1.jpg" /> 
Figure 1: Experimental Time-based Flow Chart. (A) Starting with acquired sensor plasmid DNA, E. coli vectors are transformed with DNA, selected, and amplified (Days 1-2). From concentrated E. coli LB broth, plasmid DNA is isolated and quantified (Day 3). Using purified DNA plasmids, NIH3T3 fibroblasts are transfected in a six-well format (B) and transferred onto glass-bottom dishes that are compatible with an inverted confocal microscope (Day 3). To verify the success of the transfection, cells are viewed under a widefield fluorescent microscope equipped with the appropriate filter sets (Day 4). Contingent upon a successful transfection, cells are imaged on a confocal microscope equipped with a spectral detector. Using spectral unmixing, mTFP1 and venus channels are separated during the acquisition and saved as spectrally resolved image stacks (Days 4-5). Lastly, images are preprocessed in ImageJ, and the ratio-images are computed. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54902/54902fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54902/54902fig2.jpg" /> 
Figure 2: A Representative Image of Transfected NIH3T3 Fibroblasts. (A) 10X magnification of transfected NIH3T3 fibroblasts expressing Nesprin-TS using a GFP filterset. (B) An exploded view of the bounding box from part A. Sensor expression follows the nuclear envelope and often extends into the cytoplasm. (C) Nesprin-TS and -HL Control and how they bind to the actin and SUN binding domains. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54902/54902fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54902/54902fig3.jpg" /> 
Figure 3: Spectral Fingerprinting Tutorial Using Confocal Software. Normalized emission spectra obtained from 20-pixel radii in the highlighted regions-of-interest delineated by colored crosshairs. (Arrow-1) Spectral detection mode to capture the image. (Arrow-2) &#34;Spectral Unmixing&#34; setting to add normalized spectra to the spectral database. (Arrow-3) The excitation frequency. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54902/54902fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54902/54902fig4.jpg" /> 
Figure 4: Live Spectral Unmixing and Output of Confocal Software. (A) Critical imaging parameters to reproduce live, unmixed images. (Arrow-1) The excitation frequency. (Arrow-2) Live spectral unmixing mode. (B) Nuclei image in unmixed venus channel. (C) Nuclei image in unmixed mTFP1 channel. (D) Combined image of venus and mTFP1. (Arrow-3) The nuclear membrane of the cell where Nesprin-TS is being expressed. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54902/54902fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54902/54902fig5.jpg" /> 
Figure 5: Nesprin-TS FRET ratio images in patterned and unpatterned Madin-Darby Canine Kidney (MDCK) cells. MDCK cells that stably express Nesprin-TS were imaged on 10 &#181;m-wide fibronectin lines (A) or unpatterned polydimethylsiloxane (PDMS) membranes (B). Nuclear membranes were visually masked and processed into FRET ratios. The colored scale bar represents the FRET ratio amplitude for unpatterned and patterned nuclei. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54902/54902fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/54902/54902fig6.jpg" /> 
Figure 6: Processed Ratiometric FRET Images from Nesprin-TS-Expressing Cells and Histogram Analysis of Aggregated Data. (A) A representative, normalized stacked histogram of FRET ratio images of Nesprin-TS treated with ML7 myosin inhibitor, color-coded by individual cell nuclei. (B) A representative, normalized stacked histogram of FRET ratio images of Nesprin-TS treated with Calyculin A, an activator of cell contractility. The legend correlates each analyzed nucleus to its location on the histogram. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54902/54902fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Protocol for Using FÃ¶rster Resonance Energy Transfer FRET-force Biosensors to Measure Mechanical Forces across the Nuclear LINC Complex at JoVE.com

A Protocol for Using F&#246;rster Resonance Energy Transfer (FRET)-force Biosensors to Measure Mechanical Forces across the Nuclear LINC Complex

A number of FRET-based force biosensors have recently been developed, enabling the protein-specific resolution of intracellular force. In this protocol, we demonstrate how one of these sensors, designed for the linker of the nucleoskeleton-cytoskeleton (LINC) complex protein Nesprin-2G can be used to measure actomyosin forces on the nuclear LINC complex.

The LINC complex has been hypothesized to be the critical structure that mediates the transfer of mechanical forces from the cytoskeleton to the nucleus. ...

a-protocol-for-using-frster-resonance-energy-transfer-fret-force

Nanyang Technological University

Research

JoVE Journal

Bioengineering

10.6K Views.  Virginia Commonwealth University. A number of FRET-based force biosensors have recently been developed, enabling the protein-specific resolution of intracellular force. In this protocol, we demonstrate how one of these sensors, designed for the linker of the nucleoskeleton-cytoskeleton (LINC) complex protein Nesprin-2G can be used to measure actomyosin forces on the nuclear LINC complex. 

Video: A Protocol for Using Förster Resonance Energy Transfer (FRET)-force Biosensors to Measure Mechanical Forces across the Nuclear LINC Complex

Structural Information from Single-molecule FRET Experiments Using the Fast Nano-positioning System

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. Thereby, multiple smFRET measurements are used to localize an unknown dye position inside a protein complex by means of trilateration. In order to obtain quantitative information, the Nano-Positioning System (NPS) uses probabilistic data analysis to combine structural information from X-ray crystallography with single-molecule fluorescence data to calculate not only the most probable position but the complete three-dimensional probability distribution, termed posterior, which indicates the experimental uncertainty. The concept was generalized for the analysis of smFRET networks containing numerous dye molecules. The latest version of NPS, Fast-NPS, features a new algorithm using Bayesian parameter estimation based on Markov Chain Monte Carlo sampling and parallel tempering that allows for the analysis of large smFRET networks in a comparably short time. Moreover, Fast-NPS allows the calculation of the posterior by choosing one of five different models for each dye, that account for the different spatial and orientational behavior exhibited by the dye molecules due to their local environment.
Here we present a detailed protocol for obtaining smFRET data and applying the Fast-NPS. We provide detailed instructions for the acquisition of the three input parameters of Fast-NPS: the smFRET values, as well as the quantum yield and anisotropy of the dye molecules. Recently, the NPS has been used to elucidate the architecture of an archaeal open promotor complex. This data is used to demonstrate the influence of the five different dye models on the posterior distribution.

We present the setup and experimental procedure to obtain smFRET data from large donor-acceptor networks with a TIRF microscope. The step-by-step analysis of these measurements with the Bayesian inference software Fast-NPS yields high-resolved structural information via the application of adapted dye models.

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. ...

Biochemistry

Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

The generation and detection of mechanical forces is a ubiquitous aspect of cell physiology, with direct relevance to cancer metastasis1, atherogenesis2 and wound healing3. In each of these examples, cells both exert force on their surroundings and simultaneously enzymatically remodel the extracellular matrix (ECM). The effect of forces on ECM has thus become an area of considerable interest due to its likely biological and medical importance4-7.
	Single molecule techniques such as optical trapping8, atomic force microscopy9, and magnetic tweezers10,11 allow researchers to probe the function of enzymes at a molecular level by exerting forces on individual proteins. Of these techniques, magnetic tweezers (MT) are notable for their low cost and high throughput. MT exert forces in the range of ~1-100 pN and can provide millisecond temporal resolution, qualities that are well matched to the study of enzyme mechanism at the single-molecule level12. Here we report a highly parallelizable MT assay to study the effect of force on the proteolysis of single protein molecules. We present the specific example of the proteolysis of a trimeric collagen peptide by matrix metalloproteinase 1 (MMP-1); however, this assay can be easily adapted to study other substrates and proteases.

In this article we describe the use of magnetic tweezers to study the effect of force on enzymatic proteolysis at the single molecule level in a highly parallelizable manner.

The  generation and detection of mechanical forces is a ubiquitous aspect of cell physiology,  with direct relevance to cancer metastasis1, atherogenesis2 ...

A Direct Force Probe for Measuring Mechanical Integration Between the Nucleus and the Cytoskeleton

The mechanical properties of the nucleus determine its response to mechanical forces generated in cells. Because the nucleus is molecularly continuous with the cytoskeleton, methods are needed to probe its mechanical behavior in adherent cells. Here, we discuss the direct force probe (DFP) as a tool to apply force directly to the nucleus in a living adherent cell. We attach a narrow micropipette to the nuclear surface with suction. The micropipette is translated away from the nucleus, which causes the nucleus to deform and translate. When the restoring force is equal to the suction force, the nucleus detaches and elastically relaxes. Because the suction pressure is precisely known, the force on the nuclear surface is known. This method has revealed that nano-scale forces are sufficient to deform and translate the nucleus in adherent cells, and identified cytoskeletal elements that enable the nucleus to resist forces. The DFP can be used to dissect the contributions of cellular and nuclear components to nuclear mechanical properties in living cells.

In this protocol, we describe a micropipette method to directly apply a controlled force to the nucleus in a living cell. This assay allows interrogation of nuclear mechanical properties in the living, adherent cell.

The mechanical properties of the nucleus determine its response to mechanical forces generated in cells. Because the nucleus is molecularly continuous ...

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions are likely affected by the mechanical environment in which binding and signaling take place. A recent study demonstrated that mechanical force can regulate antigen recognition by and triggering of the T-cell receptor (TCR). This was made possible by a new technology we developed and termed fluorescence biomembrane force probe (fBFP), which combines single-molecule force spectroscopy with fluorescence microscopy. Using an ultra-soft human red blood cell as the sensitive force sensor, a high-speed camera and real-time imaging tracking techniques, the fBFP is of ~1 pN (10-12 N), ~3 nm and ~0.5 msec in force, spatial and temporal resolution. With the fBFP, one can precisely measure single receptor-ligand binding kinetics under force regulation and simultaneously image binding-triggered intracellular calcium signaling on a single live cell. This new technology can be used to study other membrane receptor-ligand interaction and signaling in other cells under mechanical regulation.

We describe a technique for concurrently measuring force-regulated single receptor-ligand binding kinetics and real-time imaging of calcium signaling in a single T lymphocyte.

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions ...

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen Presenting Cells (APCs). This is despite T-cell antigen receptors (TCRs) exerting usually a moderate affinity (&#181;M range) to antigen when binding is measured in vitro1. In view of the molecular and cellular parameters contributing to T-cell antigen sensitivity, a microscopy-based methodology has been developed as a means to monitor TCR-pMHC binding in situ, as it occurs within the synapse of a live T-cell and an artificial and functionalized glass-supported planar lipid bilayer (SLB), which mimics the cell membrane of an Antigen presenting Cell (APC) 2. Measurements are based on F&#246;rster Resonance Energy Transfer (FRET) between a blue- and red-shifted fluorescent dye attached to the TCR and the pMHC. Because the efficiency of FRET is inversely proportional to the sixth power of the inter-dye distance, one can employ FRET signals to visualize synaptic TCR-pMHC binding. The sensitive of the microscopy approach supports detection of single molecule FRET events. This allows to determine the affinity and off-rate of synaptic TCR-pMHC interactions and in turn to interpolate the on-rate of binding. Analogous assays could be applied to measure other receptor-ligand interactions in their native environment.

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

This manuscript describes how to conduct (single molecule) F&#246;rster Resonance Energy Transfer (FRET)- based assays to measure the binding dynamics between T-cell antigen receptor (TCR) and antigenic peptide-loaded MHC molecules as they occur within the immunological synapse of a T-cell in contact with a functionalized planar supported lipid bilayer.

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen ...

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Developmental Biology

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.

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 ...

Automated Two-dimensional Spatiotemporal Analysis of Mobile Single-molecule FRET Probes

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. It has been used in a wide range of biophysical and molecular biological experiments, including the measurement of molecular forces, characterization of conformational dynamics of biomolecules, observation of intracellular colocalization of proteins, and determination of receptor-ligand interaction times. In a widefield microscopy configuration, experiments are typically performed using surface-immobilized probes. Here, a method combining single-molecule tracking with alternating excitation (ALEX) smFRET experiments is presented, permitting the acquisition of smFRET time traces of surface-bound, yet mobile probes in plasma membranes or glass-supported lipid bilayers. For the analysis of recorded data, an automated, open-source software collection was developed supporting (i) the localization of fluorescent signals, (ii) single-particle tracking, (iii) determination of FRET-related quantities including correction factors, (iv) stringent verification of smFRET traces, and (v) intuitive presentation of the results. The generated data can conveniently be used as input for further exploration via specialized software, e.g., for the assessment of the diffusional behavior of probes or the investigation of FRET transitions.

This article presents a method for spatiotemporal analysis of mobile, single-molecule F&#246;rster resonance energy transfer (smFRET)-based probes using widefield fluorescence microscopy. The newly developed software toolkit allows the determination of smFRET time traces of moving probes, including the correct FRET efficiency and the molecular positions, as functions of time.

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. ...

FLIM-FRET Measurements of Protein-Protein Interactions in Live Bacteria.

Protein-protein interactions (PPIs) control various key processes in cells. Fluorescence lifetime imaging microscopy (FLIM) combined with F&#246;rster resonance energy transfer (FRET) provide accurate information about PPIs in live cells. FLIM-FRET relies on measuring the fluorescence lifetime decay of a FRET donor at each pixel of the FLIM image, providing quantitative and accurate information about PPIs and their spatial cellular organizations. We propose here a detailed protocol for FLIM-FRET measurements that we applied to monitor PPIs in live Pseudomonas aeruginosa in the particular case of two interacting proteins expressed with highly different copy numbers to demonstrate the quality and robustness of the technique at revealing critical features of PPIs. This protocol describes in detail all the necessary steps for PPI characterization - starting from bacterial mutant constructions up to the final analysis using recently developed tools providing advanced visualization possibilities for a straightforward interpretation of complex FLIM-FRET data.

We describe here a protocol to characterize protein-protein interactions between two highly-differently expressed proteins in live Pseudomonas aeruginosa using FLIM-FRET measurements. The protocol includes bacteria strain constructions, bacteria immobilization, imaging and post-imaging data analysis routines.

Protein-protein interactions (PPIs) control various key processes in cells. Fluorescence lifetime imaging microscopy (FLIM) combined with F&#246;rster ...

A Protocol for Using Förster Resonance Energy Transfer (FRET)-force Biosensors to Measure Mechanical Forces across the Nuclear LINC Complex

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