This project is funded by UF Gatorade Award Start-up Package (X. T.), the UFHCC Pilot Award (X. T. and Dr. Dietmar Siemann), UF Opportunity Seed Fund (X. T.), and UFHCC University Scholars Program (H. Y. Wang). We sincerely appreciate the intellectual discussions with and the technical support from Dr. Jonathan Licht (UFHCC), Dr. Rolf Renne (UFHCC), Dr. Christopher Vulpe (UFHCC), Dr. Blanka Sharma (BME), Dr. Mark Sheplak (MAE &#38; ECE), Dr. Daniel Ferris (BME), Dr. Malisa Sarntinoranont (MAE), Dr. Ashok Kumar (MAE), Dr. Benjamin Keselowsky (BME), Dr. Brent Gila (RSC), Dr. Philip Feng (ECE), Dr. Gregory A. Hudalla (BME), Dr. Steven Ghivizzani (OSSM), Dr. Yenisel Cruz-Almeida (CDBS), Dr. Roger Fillingim (CD-BS), Dr. Robert Caudle (OMS), Dr. John Neubert (DN-OR), Dr. Justin Hiliard (Neurosurgery), Dr. Tian He (Harvard University), Dr. Youhua Tan (Hong Kong Polytechnic University), Dr. Jessie L-S Au (Institute of Quantitative Systems Pharmacology), Dr. David Hahn (University of Arizona), and Support Team of Nikon (Drs. Jose Serrano-Velez, Larry Kordon, and Jon Ekman). We are deeply grateful for the effective support from all members of Tang&#39;s, Yamaguchi&#39;s, Sharma&#39;s, Au&#39;s, Siemann&#39;s, and Guan&#39;s research laboratories and all staff members of the UF MAE Department.

1. Maintenance of CRISPR/Cas9-engineered B2B cells
<ol>
	<li>Culture B2B cells in a T25 flask with RPMI-1640 supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin.</li>
	<li>Maintain the B2B cells in a humidified incubator at 37 &#176;C with 5% CO2.</li>
	<li>Subculture the B2B cells when the confluency reaches 70% to 80%.</li>
	<li>Store the B2B cell line in RPMI-1640 culture medium with 10% (v/v) DMSO in a -80 &#176;C freezer.</li>
	<li>Use the B2B cells with a passage number less than 10 in the experiments.</li>
</ol>
2. Cell culture
<ol>
	<li>Seed the cells onto a glass-bottom Petri dish.
	<ol>
		<li>Move the flask that contains B2B cells inside from the incubator to the biosafety cabinet.</li>
		<li>Remove the culture medium in the flask using an aspirating pipette with a vacuum pump connected.</li>
		<li>Wash the flask with 2 mL of phosphate-buffered saline (PBS).&#8239;</li>
		<li>Remove PBS using the aspirating pipette.</li>
		<li>Add 0.5 mL of 0.05% Trypsin solution to detach cells from the bottom of the flask substrate.</li>
		<li>Put the flask in the incubator for 5 min.</li>
		<li>Move the flask to the biosafety cabinet. Add 5 mL of new culture medium into the flask and pipette the solution up and down.</li>
		<li>Deposit 50 &#181;L of the medium with cells (300 cells/&#181;L) onto the glass-bottom Petri dish. Add 2 mL of culture medium into the Petri dish.</li>
		<li>Place the Petri dish into the incubator. Wait for 12 h for the cells to attach.</li>
	</ol>
	</li>
	<li>Culture the cells with magnetic microbeads.
	<ol>
		<li>Weigh 0.2 g of 7 &#181;m mean diameter carbonyl iron microbeads (hereafter called 7 &#181;m microbeads, see the Table of Materials).</li>
		<li>Use a pipette to suspend the microbeads in 1 mL of RPMI-1640 culture medium.</li>
		<li>Take the Petri dish with B2B cells to the biosafety cabinet.</li>
		<li>Add 200 &#181;L of the medium containing microbeads into the Petri dish. 
		NOTE: Add the medium quickly to avoid precipitation of the microbeads.</li>
		<li>Put the Petri dish back in the incubator until microbeads are internalized by the cells. Check the internalization every 6 h to determine the optimal time for internalization for different cell lines.</li>
		<li>To check the internalization, perform confocal fluorescence imaging to visualize the microbead, nuclear, and cell boundary. If the microbead is internalized by the cell, it will be within the cell boundary.</li>
	</ol>
	</li>
</ol>
3. Visualization of nucleus
<ol>
	<li>Warm 1.5 mL of the culture medium in the incubator for 15 min.</li>
	<li>Turn off the light of the biosafety cabinet. Take the Petri dish that contains the cell, warmed culture medium, nuclear stain, and Verapamil HCl into the biosafety cabinet. 
	NOTE: Nuclear staining components are sensitive to light. Avoid exposure to light during operation.</li>
	<li>Dilute 1000x nuclear stain by DMSO to 100x.</li>
	<li>Dilute 100 mM Verapamil HCl by DMSO to 10 mM.</li>
	<li>Add 15 &#181;L of 100x nuclear stain and 15 &#181;L of 10 mM Verapamil HCl to 1.5 mL of culture medium. Mix well by pipetting up and down.</li>
	<li>Remove the culture medium from the Petri dish. Add the culture medium containing nuclear staining into the Petri dish.</li>
	<li>Put the cells back in the incubator for over 2 h.</li>
</ol>
4. Preparation of the magnetic force application hardware
<ol>
	<li>3D print all parts using acrylonitrile butadiene styrene (ABS) and assemble them following the CAD design (Figure 1A). The CAD design is included in the Table of Materials.</li>
	<li>Use double-sided tape to attach the magnet to the magnet-moving device (Figure 1A).</li>
	<li>Set the magnet-moving device next to the microscope stage. Use the three knobs to adjust the spatial location of the magnet until it can move above the Petri dish between 13 mm and 120 mm. 
	NOTE: Ensure the upper limit of the distance between the magnet and Petri dish is as large as possible to avoid unwanted force application on the magnetic microbeads. 120 mm is the maximum value in this experimental setup. Ensure that the magnet does not interfere with microscope parts, including objectives and motorized stages.</li>
	<li>Set the magnet to the highest z-position (at 120 mm).</li>
</ol>
5. Force application and live-cell imaging
<ol>
	<li>Set up of the environment chamber for long-term imaging
	<ol>
		<li>Apply 75% ethanol solution to thoroughly sterilize and clean the environment chamber.</li>
		<li>Place the environment chamber onto the motorized stage of the inverted microscope.</li>
		<li>Open the CO2 tank and set the CO2 inflow rate to 160 mL/min.</li>
		<li>Adjust the temperature of the chamber to 44 &#176;C (Top), 42 &#176;C (Bath), and 40 &#176;C (Stage).</li>
		<li>Add 20 mL of purified water into the bath of the environment chamber to maintain 90% humidity.</li>
		<li>Take out the glass-bottom Petri dish that contains target cells from the tissue culture incubator and place it into the chamber.</li>
		<li>Apply the metal clamp of the environment chamber to fix the Petri dish position. 
		NOTE: The Petri dish must be clamped tightly in the chamber because the magnetic force may move the dish if it is not clamped.</li>
		<li>Close the lid of the chamber.&#8239;</li>
	</ol>
	</li>
	<li>Optimization of imaging parameters
	<ol>
		<li>Optimize the pinhole size: The pinhole blocks the out-of-focus photons. A larger pinhole size yields more out-of-focus photons but a brighter image. A smaller pinhole size yields a more focused and dimmer image. Make sure to optimize the pinhole size to get in-focus confocal images with the appropriate signal-to-noise ratio.</li>
		<li>Optimize the laser intensity: The laser intensity determines the intensity of excitation and thus emission light. The low laser intensity gives a low signal-to-noise ratio. Too high a laser intensity will cause photobleaching. Adjust the laser intensity accordingly.</li>
		<li>Optimize the step size and steps: Steps and step size determine how many images will take in a Z-stack. Smaller step sizes and more steps will increase the Z-stack resolution but will also increase photobleaching. In this experiment, 1 &#181;m step size was used for the cells with ~15 &#181;m cell height.</li>
		<li>Optimize the exposure time: The exposure time determines how long the cell will be exposed to the excitation laser. A low exposure time will decrease the signal-to-noise ratio. A high exposure time will cause photobleaching. An exposure time of 1 frame per 4 s was used in this experiment.</li>
		<li>Optimization of imaging parameters: Change one of the four parameters iteratively and keep the other parameters consistent. Each time, measure the YAP N/C ratio of each image and compare the YAP N/C ratio change to determine the photobleaching level. Repeat the optimization process until achieving a balance between the signal-to-noise ratio, imaging speed, and photobleaching.</li>
		<li>Define the imaging configurations using the optimized imaging parameters for faster imaging settings during the experiments. 
		NOTE: Configurations used in this study are described in section 5.3 of imaging parameters. To optimize imaging parameters of configurations in section 5.3, use the same method as in step 5.2.5.</li>
	</ol>
	</li>
	<li>Small force application and confocal imaging 
	NOTE: Nikon Ti2-E microscope was used for imaging in this study, and detailed steps for image acquisition are given below.
	<ol>
		<li>Open the inverted microscope. Open the software application Elements.</li>
		<li>Define configuration magnetic_find. Check only FITC channel. Set PMT HV = 70, Offset = 0, Laser intensity = 10. Set the scanning speed to 1 frame per 2 s by clicking the 1/2 button. Set pinhole size to 1.2 AU by clicking the 1.2 A.U. button. This configuration will be used in step 5.3.5.</li>
		<li>Define configuration magnetic_YAP_Nucleus. Check FITC channel. Set PMT HV = 70, Offset = 0, Laser intensity = 10. Set the scanning speed to 1 frame per 4 s by clicking the 1/2 button. Set pinhole size to 1.2 AU by clicking the 1.2 A.U. button. To image the nucleus boundary and nuclear stain intensity, check Cy5 channel. Set PMT HV = 70, Offset = 0, Laser intensity = 10. The pinhole size is optimized for 3D YAP imaging. Do not click the 1.2 A.U. button again after checking the Cy5 channel. This configuration will be used in step 5.3.7.</li>
		<li>Turn on DIA through Elements if necessary. Open SpinView, use a bright-field, and adjust the focus of the object to get a clear in-focus image of cells. Use a 10x objective to find appropriate multiple single cells in three conditions: with a single microbead inside, with multiple microbeads inside, and without any microbead inside. Switch to 40x objective. Name this position with the appropriate position number.</li>
		<li>Open Elements. Click on magnetic_find. Click the Remove Interlock button.</li>
		<li>Click Scan and adjust the Z-position of the focal plane. Click the Top and Bottom buttons to set the lower and upper limit for the Z-stack of the selected cells. Stop scanning by clicking Scan again.</li>
		<li>Switch to magnetic_YAP_Nucleus configuration. Set file name as before_small_force.nd2. Click on the Run button with the recorded Z-stack.</li>
		<li>Switch to the right light path and turn on DIA. Open SpinView and click on the Recording button. Meanwhile, spin the knob of the magnet-moving device to move the magnet down to 46 mm above the Petri dish bottom. Save bright-field image sequence or video. Check the video to confirm microbeads show displacement induced by magnetic force.</li>
		<li>Repeat steps 5.3.5-5.3.7; set the file name to after_small_force.nd2.</li>
		<li>Switch to the right light path and turn on DIA. Next, open SpinView and click on the Recording button. Meanwhile, spin the knob of the magnet-moving device to move the magnet up to 120 mm above the Petri dish bottom. Save bright-field image sequence or video.</li>
		<li>Repeat steps 5.3.5-5.3.7 and set the file name to before_large_force.nd2.</li>
	</ol>
	</li>
	<li>Large force application and confocal imaging
	<ol>
		<li>Remove the lid of the environment chamber to allow the magnet to reach 13 mm above the Petri dish bottom.</li>
		<li>Switch to the right light path and turn on DIA. Open SpinView and click on the Recording button. Meanwhile, spin the knob of the magnet-moving device to move the magnet down to 13 mm above the Petri dish bottom. Save bright-field image sequence or video. Check the video to confirm microbeads show displacement induced by magnetic force.</li>
		<li>Repeat steps 5.3.5-5.3.7 and set the file name to after_large_force.nd2.</li>
		<li>Switch to the right light path and turn on DIA. Next, open SpinView and click on the Recording button. Meanwhile, spin the knob of the magnet-moving device to move the magnet up to 120 mm above the Petri dish bottom. Save bright-field image sequence or video.</li>
		<li>Repeat steps 5.3.5-5.3.7; set the file name to retract_large_force.nd2.</li>
		<li>Close the lid of the environment chamber.</li>
	</ol>
	</li>
	<li>Repeat steps 5.2 and 5.3 for multiple fields of view to obtain more data if needed.</li>
</ol>
6. Image processing and data analysis
<ol>
	<li>Quantification of YAP N/C ratio
	<ol>
		<li>Open Fiji ImageJ. Open the .nd2 images taken in step 5.</li>
		<li>Click on Analyze &#62; Set Measurements. Check Area, Integrated Density, Mean Grey Value, and Shape Descriptors.</li>
		<li>Use the Cy5 channel to identify the nucleus. Click on Freehand Selections to use the free-selection tool to outline the nucleus. Also, check the automatic nuclear mask macro in ImageJ (see the Table of Materials).</li>
		<li>Click Analyze &#62; Measure in the FITC channel. The measured value of the Mean is the average nuclear YAP intensity DN.</li>
		<li>Use the Cy5 channel to identify the nucleus. Use the FITC channel to identify the cell. Click Freehand Selections to use the free-selection tool to select a region of interest within the cytoplasm and avoid the magnetic microbead. This region of interest must not include the nucleus.</li>
		<li>Click Analyze &#62; Measure in the FITC channel. The measured value of the Mean is the average cytoplasmic YAP intensity DC.</li>
		<li>Calculate the YAP N/C ratio = DN / DC.</li>
	</ol>
	</li>
	<li>Quantification of nuclear shape and normalized nuclear stain intensity
	<ol>
		<li>Open Fiji ImageJ. Open the .nd2 images taken in step 5.</li>
		<li>Click on Analyze &#62; Set Measurements. Check Area, Integrated Density, Mean Grey Value, and Shape Descriptors.</li>
		<li>Use the Cy5 channel to identify the nucleus. Click on Freehand Selections to use the free-selection tool to outline the nucleus.</li>
		<li>Click on Analyze &#62; Measure in Cy5 channel. The measured value of the Mean is the nuclear stain intensity. The measured value of Circ. is nuclear circularity.</li>
		<li>To compare the nuclear stain intensity at different force state, all nuclear stain intensity is divided by the nuclear stain intensity in &#34;before_small_force.nd2&#34; to generate the normalized nuclear stain intensity.</li>
	</ol>
	</li>
</ol>

There are no conflicts of interest to declare.

Internalization of magnetic microbeads (section 2.2) is critical because extracellular microbeads cannot apply force directly to the nucleus. Force application and imaging (section 5.3)&#160;are&#160;critical steps in this experiment, and the force needed to deform the nucleus and induce meaningful biological consequences might be sample-dependent. The force magnitude in this experiment (0.8 nN and 1.4 nN) can be further increased to trigger nuclear mechano-sensing in less sensitive cells.
To ...

This study aims to develop and apply a new technique to elucidate nucleus mechanobiology by combining the magnetic actuators that apply mechanical force directly on the cell nucleus and the confocal fluorescence microscopy that simultaneously images the structural and functional subcellular changes. Cells sense extracellular biophysical signals including tissue stiffness1,2,3,4, interstitial fluid pressure and shear stress5,6,7, surface topology/geometry8,9,10,11,12, and tension/compression stress13,14,15,16. Biophysical signals are converted into biochemical signals and trigger potential downstream changes of gene expression and cell behaviors-a process known as mechanotransduction17,18,19,20,21,22,23,24,25,26,27. To study mechanotransduction processes, a myriad of techniques have been developed to apply mechanical force on the cells, such as atomic force microscopy28, cell stretching device29, bio-MEMS (micro-electromechanical systems) force sensor15,30,31, shear rheology32, and Stereo Vision System33. A recent review summarizes the approaches to apply extracellular mechanical cues and interfere with mechanosensing34. To date, most of these methods apply force on the cell plasma membrane, and cells directly receive these extracellular biophysical signals via membrane receptors such as integrin, cadherin, ion channels, and G-Protein-coupled receptors. Subsequently, they transmit the signal to the intracellular cytoskeleton and nucleus. For example, using yes-associated protein (YAP) translocation as an indicator of mechano-sensing, cells are shown to sense the mechanical signals of substrate stiffness and extracellular tension from the cell membrane and transmit them through the cytoskeleton into the nucleus to induce YAP cytoplasm-to-nucleus translocation28,35.
Recent evidence suggests that the cell nucleus itself is an independent mechano-sensor8,36,37. This is proven by experiments performed on the isolated nucleus harvested from cells, where it was revealed that nuclei adaptively change their stiffness in response to the mechanical force directly applied on them36. During many physiological conditions, nuclei in both tumor and healthy cells sense extracellular biophysical signals and change their mechanical properties and assemblies38,39,40. For example, upon extravasation, the nuclear stiffness of tumor cells decreases and maintains softness for over 24 h38. During migration through confined interstitial space, the nuclei of tumor cells frequently lose and recover their structural integrity39. However, the way in which the nucleus senses the biophysical signal is unknown, although several nuclear-envelope proteins and families of proteins have been found to be involved, such as Lamin A/C and linker of nucleoskeleton and cytoskeleton (LINC) complex38,41. Hence, new non-invasive methods that can directly apply force to the nucleus will decouple the effect of force transmission from the cell-plasma membrane and cytoskeleton, and will help elucidate the previously inaccessible molecular mechanisms of nuclear mechano-sensing.
Research that employed optical tweezers to manipulate organelles42 and microbeads injected into cells43 showed the technological capability of directly applying force on the nucleus. However, the optical-tweezer technique has several limitations: (1) low throughput-optical tweezers often only manipulate one cell or microbead at a time; and (2) potential photodamage and temperature artifact-deformation of nuclear requires tens of pN36, and the corresponding necessary laser power is about 10 mW per pN44,45. Such laser intensity is sufficient to trigger photodamage in the cells and perturb cell functions during the experiment46.
Magnetic force applied through microbeads within living cells shows the potential to directly apply force on the nucleus and overcomes the limitations of optical tweezers. Once microbeads are delivered into the cytoplasm, a magnetic field can exert a magnetic force on multiple microbeads simultaneously in a high-throughput manner. The magnetic field does not influence cell functions47,&#160;but generates force from pN to nN, which is enough to induce nuclear deformation36,48,49.&#160;To date, manipulation of magnetic microbeads has been applied on cell plasma membrane48, inside the cytoplasm50, on F-actin51, inside the nucleus47, and on the isolated nucleus36. However, magnetic manipulation of microbeads has never been used to apply direct mechanical force on the nuclear envelope to study mechanotransduction in the nucleus.
In this paper, a simple technique is developed to non-invasively deliver magnetic microbeads into the cytoplasm and use these microbeads to apply mechanical force on the nucleus (Figure 1). Here, CRISPR/Cas9-engineered human normal B2B cell lines that endogenously express mNeonGreen21-10/11-tagged YAP are used to validate the method. YAP is a mechano-sensitive protein, and the translocation of YAP is regulated by nuclear mechano-sensing14,28. The CRISPR/Cas9-regulated knock-in approach was chosen to tag endogenous YAP with a fluorescent protein (FP) mNeonGreen21-10/11. Although CRISPR editing is known to have incomplete efficiency and off-target effect, the protocols in previous publications integrated fluorescence sorting to select for correct open reading frame insertion52,53,54. With this additional layer of selection, no off-target tagging event was observed in 20+ cell lines previously generated52,53,54,55. This is a split fluorescent protein construct, but in principle, any expressible fluorescent tag could be usable. This labeling approach is superior to transgene or antibody methods. First, unlike the transgene expression, the tagged protein maintains single-copy gene dosage and expresses in the physiological context of the native gene regulatory network, limiting deviations in protein concentration, localization, and interaction. The tagging method used in this study achieves over an order-of-magnitude higher throughput and efficiency than full FP tagging. It also avoids challenges associated with immunofluorescence due to fixation artifacts and the limited availability of high-quality, high-specificity antibodies. Second, the approach used in this paper does minimum perturbation to the cell physiology and enables the real-time revelation of all endogenous YAPs authentically. In contrast, other common transgene methods often lead to overexpression of YAP. The resulting artificial distribution can potentially cause cytotoxicity and affect mechano-sensing of cells56,57,58.
This study presents a protocol to directly apply force on the nucleus through magnetic microbeads delivered into the cytoplasm and to conduct simultaneous live-cell fluorescent imaging. In summary, the protocols presented here demonstrate how to (1) deliver magnetic microbeads into the cell while outside the nucleus, (2) manipulate the microbeads to apply magnetic force on the nucleus, (3) perform confocal fluorescent imaging of the cells during manipulation, and (4) quantitatively analyze the YAP nuclear/cytoplasm (N/C) ratio throughout the force application process. The results suggest that (1) through endocytosis, magnetic microbeads can be non-invasively delivered into the cytoplasm of B2B cells within 7 h (Figure 2 and Figure 3); and (2) quantified magnetic force directly applied on the nucleus (Figure 4, Figure 5, and Figure 6) alone can trigger diverse changes of YAP N/C ratio in CRISPR/Cas9-engineered B2B cells (Figure 7 and Figure 8).

<table><tbody><tr><td>0.05 % Trypsin</td><td>Corning</td><td>25-051-CI</td><td></td></tr><tr><td>25 cm&lt;sup&gt;2&lt;/sup&gt; flask</td><td>Corning</td><td>156340</td><td></td></tr><tr><td>7-&amp;micro;m mean diameter carbonyl iron microbeads</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>A1R confocal system</td><td>Nikon</td><td></td><td></td></tr><tr><td>Carbonyl Iron Powder CM</td><td>BASF</td><td>30042253</td><td>Magnetic microbead</td></tr><tr><td>Culture medium (RPMI-1640)</td><td>Gibco</td><td>11875093</td><td></td></tr><tr><td>Desktop Computer</td><td>Dell</td><td></td><td>with Windows 10 operating system</td></tr><tr><td>Environmental chamber TIZB</td><td>Tokai Hit</td><td>TIZB</td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Gibco</td><td>26140</td><td></td></tr><tr><td>Fiji ImageJ</td><td>National Institutes of Health and the Laboratory for Optical and Computational Instrumentation</td><td></td><td></td></tr><tr><td>Glass-bottom petri dish</td><td>MatTek</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>Magnet</td><td>K&amp;amp;J Magnetics, Inc.</td><td>D99-N52</td><td></td></tr><tr><td>Monochrome&amp;nbsp;Camera</td><td>FLIR</td><td>BFS-U3-70S7M-C</td><td></td></tr><tr><td>NIS-Elements software platform</td><td>Nikon</td><td></td><td>software platform</td></tr><tr><td>Nucleus mask ImageJ macro</td><td></td><td></td><td>https://github.com/KOLIUG/Nuclear mask</td></tr><tr><td>NucSpot Live 650</td><td>Biotium</td><td>#40082</td><td>Nuclear stain</td></tr><tr><td>Penicillin-streptomycin</td><td>Gibco</td><td>15140122</td><td></td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Gibco</td><td>10010023</td><td></td></tr><tr><td>Ti2-E inverted microscope</td><td>Nikon</td><td></td><td></td></tr><tr><td>XYZ mover (CAD files)</td><td></td><td></td><td>https://github.com/KOLIUG/XYZ-mover</td></tr></tbody></table>

combining 3d magnetic force actuator and multi-functional fluorescence imaging to study nucleus mechanobiology

A fundamental question in mechanobiology is how living cells sense extracellular mechanical stimuli in the context of cell physiology and pathology. The cellular mechano-sensation of extracellular mechanical stimuli is believed to be through the membrane receptors, the associated protein complex, and the cytoskeleton. Recent advances in mechanobiology demonstrate that the cell nucleus in cytoplasm itself can independently sense mechanical stimuli simultaneously. However, a mechanistic understanding of how the cell nucleus senses, transduces, and responds to mechanical stimuli is lacking, mainly because of the technical challenges in accessing and quantifying the nucleus mechanics by conventional tools. This paper describes the design, fabrication, and implementation of a new magnetic force actuator that applies precise and non-invasive 3D mechanical stimuli to directly deform the cell nucleus. Using CRISPR/Cas9-engineered cells, this study demonstrates that this tool, combined with high-resolution confocal fluorescent imaging, enables the revelation of the real-time dynamics of a mechano-sensitive yes-associated protein (YAP) in single cells as a function of nucleus deformation. This simple method has the potential to bridge the current technology gap in the mechanobiology community and provide answers to the knowledge gap that exists in the relation between nucleus mechanotransduction and cell function.

Design of a magnet-moving device and application of magnetic force 
To apply force on the nucleus through the magnetic microbeads, a magnet-moving device was designed and built to control the spatial position of the magnet. The magnet-moving device contains a central frame, three knobs, and rails to move the attached magnet in x, y, and z directions independently at the spatial resolution of 1.59 mm per cycle (Figure 1A). Once the magnet is moved close to the 7 &#181;m ...

Watch this Scientific Journal Video about Combining 3D Magnetic Force Actuator and Multi-Functional Fluorescence Imaging to Study Nucleus Mechanobiology at JoVE.com

Combining 3D Magnetic Force Actuator and Multi-Functional Fluorescence Imaging to Study Nucleus Mechanobiology

This study presents a new protocol to directly apply mechanical force on the cell nucleus through magnetic microbeads delivered into the cytoplasm and to conduct simultaneous live-cell fluorescent imaging.

A fundamental question in mechanobiology is how living cells sense extracellular mechanical stimuli in the context of cell physiology and pathology. The ...

combining-3d-magnetic-force-actuator-multi-functional-fluorescence

Research

JoVE Journal

Biology

2.3K Views.  University of Florida. Bij deze methode wordt de kracht direct op de kern uitgeoefend. Dit ontkoppelt het krachtoverdrachtseffect van het celplasmamembraan en cytoskelet en onthult de moleculaire mechanismen van nucleaire mechanosensing. De kracht wordt op een niet-invasieve manier uitgeoefend in de levende cellen.In vergelijking met optische pincetten hebben het magnetisch veld en de magnetische kracht geen invloed op de celfuncties en hebben ze een hogere doorvoer. Begin met het kweken van de cellen met ma...

Video: Combining 3D Magnetic Force Actuator and Multi-Functional Fluorescence Imaging to Study Nucleus Mechanobiology

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 ...

Bioengineering

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 ...

Biochemistry

Remote Magnetic Actuation of Micrometric Probes for in situ 3D Mapping of Bacterial Biofilm Physical Properties

Bacterial adhesion and growth on interfaces lead to the formation of three-dimensional heterogeneous structures so-called biofilms. The cells dwelling in these structures are held together by physical interactions mediated by a network of extracellular polymeric substances. Bacterial biofilms impact many human activities and the understanding of their properties is crucial for a better control of their development &#8212; maintenance or eradication &#8212; depending on their adverse or beneficial outcome. This paper describes a novel methodology aiming to measure in situ the local physical properties of the biofilm that had been, until now, examined only from a macroscopic and homogeneous material perspective. The experiment described here&#160;involves introducing magnetic particles into a growing biofilm to seed local probes that can be remotely actuated without disturbing the structural properties of the biofilm. Dedicated magnetic tweezers were developed to exert a defined force on each particle embedded in the biofilm. The setup is mounted on the stage of a microscope to enable the&#160;recording of time-lapse images of the particle-pulling period. The particle trajectories are then extracted from the pulling sequence and the local viscoelastic parameters are derived from each particle displacement curve, thereby providing the 3D-spatial distribution of the parameters. Gaining insights into the biofilm mechanical profile is essential from an engineer&#39;s point of view for biofilm control purposes but also from a fundamental perspective to clarify the relationship between the architectural properties and the specific biology of these structures.

Remote Magnetic Actuation of Micrometric Probes for in situ 3D Mapping of Bacterial Biofilm Physical Properties

This paper shows an original methodology based on the remote actuation of magnetic particles seeded in a bacterial biofilm and the development of dedicated magnetic tweezers to measure in situ the local mechanical properties of the complex living material built by micro-organisms at interfaces.

Bacterial adhesion and growth on interfaces lead to the formation of three-dimensional heterogeneous structures so-called biofilms. The cells dwelling in ...

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 ...

Magnetic Tweezers in a Microplate Format

Mechanobiology describes how the physical forces and mechanical properties of biological material contribute to physiology and disease. Typically, these approaches are limited single-molecule methods, which restricts their availability. To address this need, a microplate assay was developed that enables mechanical manipulation while performing standard biochemical assays. This is achieved using magnets incorporated into a microplate lid to create multiple magnetic tweezers. In this format, force is exerted across biomolecules connected to paramagnetic beads, equivalent to a typical magnetic tweezer. The study demonstrates the application of this tool with FRET-based assays to monitor protein conformations. However, this approach is widely applicable to different biological systems ranging from measuring enzymatic activity through to the activation of signaling pathways in live cells.

Here, we describe the use of a novel microplate assay to enable mechanical manipulation of biomolecules while performing ensemble biochemical assays. This is achieved using a microplate lid modified with magnets to create multiple static magnetic tweezers across the microplate.

Mechanobiology describes how the physical forces and mechanical properties of biological material contribute to physiology and disease. Typically, these ...

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.

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. ...

Live Cell Response to Mechanical Stimulation Studied by Integrated Optical and Atomic Force Microscopy

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to investigate cells behavior at sub-cellular level with high spatial and temporal resolution was developed. Thus, an atomic force microscope (AFM) was integrated with total internal reflection fluorescence (TIRF) microscopy and fast-spinning disk (FSD) confocal microscopy. The integrated system is broadly applicable across a wide range of molecular dynamic studies in any adherent live cells, allowing direct optical imaging of cell responses to mechanical stimulation in real-time. Significant rearrangement of the actin filaments and focal adhesions was shown due to local mechanical stimulation at the apical cell surface that induced changes into the cellular structure throughout the cell body. These innovative techniques will provide new information for understanding live cell restructuring and dynamics in response to mechanical force. A detailed protocol and a representative data set that show live cell response to mechanical stimulation are presented.

This paper aims to instruct the reader in the operation of an integrated atomic force-optical imaging microscope for mechanical stimulation of live cells in culture. A step-by-step protocol is presented. A representative data set that shows live cell response to mechanical stimulation is presented.

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to ...

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

High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the effect of applied extracellular tensile strain on cell populations in vitro via biochemical assays, a device has previously been designed which can be fabricated simply and is small enough to fit inside tissue culture incubators, as well as on top of microscope stages. However, the previous design of the polydimethylsiloxane substratum did not allow high-resolution subcellular imaging via oil-immersion objectives. This work describes a redesigned geometry of the polydimethylsiloxane substratum and a customized imaging setup that together can facilitate high-resolution subcellular imaging of live cells while under applied strain. This substratum can be used with the same, earlier designed device and, hence, has the same advantages as listed above, in addition to allowing high-resolution optical imaging. The design of the polydimethylsiloxane substratum can be improved by incorporating a grid which will facilitate tracking the same cell before and after the application of strain. Representative results demonstrate high-resolution time-lapse imaging of fluorescently labeled nuclei within strained cells captured using the method described here. These nuclear dynamics data give insights into the mechanism by which applied tensile strain promotes differentiation of oligodendrocyte progenitor cells.&#160;

Using a previously designed device to apply mechanical strain to adherent cells, this paper describes a redesigned substratum geometry and a customized apparatus for high-resolution single-cell imaging of strained cells with a 100x oil immersion objective.

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the ...

Biophysical Assays to Probe the Mechanical Properties of the Interphase Cell Nucleus: Substrate Strain Application and Microneedle Manipulation

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the physical properties of the nucleus contribute significantly to the overall biomechanical behavior of cells under physiological and pathological conditions. For example, in migrating neutrophils and invading cancer cells, nuclear stiffness can pose a major obstacle during extravasation or passage through narrow spaces within tissues.1 On the other hand, the nucleus of cells in mechanically active tissue such as muscle requires sufficient structural support to withstand repetitive mechanical stress. Importantly, the nucleus is tightly integrated into the cellular architecture; it is physically connected to the surrounding cytoskeleton, which is a critical requirement for the intracellular movement and positioning of the nucleus, for example, in polarized cells, synaptic nuclei at neuromuscular junctions, or in migrating cells.2 Not surprisingly, mutations in nuclear envelope proteins such as lamins and nesprins, which play a critical role in determining nuclear stiffness and nucleo-cytoskeletal coupling, have been shown recently to result in a number of human diseases, including Emery-Dreifuss muscular dystrophy, limb-girdle muscular dystrophy, and dilated cardiomyopathy.3 To investigate the biophysical function of diverse nuclear envelope proteins and the effect of specific mutations, we have developed experimental methods to study the physical properties of the nucleus in single, living cells subjected to global or localized mechanical perturbation. Measuring induced nuclear deformations in response to precisely applied substrate strain application yields important information on the deformability of the nucleus and allows quantitative comparison between different mutations or cell lines deficient for specific nuclear envelope proteins. Localized cytoskeletal strain application with a microneedle is used to complement this assay and can yield additional information on intracellular force transmission between the nucleus and the cytoskeleton. Studying nuclear mechanics in intact living cells preserves the normal intracellular architecture and avoids potential artifacts that can arise when working with isolated nuclei. Furthermore, substrate strain application presents a good model for the physiological stress experienced by cells in muscle or other tissues (e.g., vascular smooth muscle cells exposed to vessel strain). Lastly, while these tools have been developed primarily to study nuclear mechanics, they can also be applied to investigate the function of cytoskeletal proteins and mechanotransduction signaling.

We present two independent, microscope-based tools to measure the induced nuclear and cytoskeletal deformations in single, living adherent cells in response to global or localized strain application. These techniques are used to determine nuclear stiffness (i.e., deformability) and to probe intracellular force transmission between the nucleus and the cytoskeleton.

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the ...

Combining 3D Magnetic Force Actuator and Multi-Functional Fluorescence Imaging to Study Nucleus Mechanobiology

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