MK acknowledges financial support from the Spanish Ministry of Economy and Competitiveness through the Plan Nacional (PGC2018-097882-A-I00), FEDER (EQC2018-005048-P), Severo Ochoa program for Centres of Excellence in R&#38;D (CEX2019-000910-S; RYC-2016-21062), from Fundaci&#243; Privada Cellex, Fundaci&#243; Mir-Puig, and from Generalitat de Catalunya through the CERCA and Research program (2017 SGR 1012), in addition to funding through ERC (MechanoSystems) and HFSP (CDA00023/2018). V.R. acknowledges support from the Spanish Ministry of Science and Innovation to the EMBL partnership, the Centro de Excelencia Severo Ochoa, MINECO&#39;s Plan Nacional (BFU2017-86296-P, PID2020-117011GB-I00) and Generalitat de Catalunya (CERCA). V.V. acknowledges support from the ICFOstepstone PhD Programme funded by the European Union&#39;s Horizon 2020 research and innovation program under Marie Sk&#322;odowska-Curie grant agreement 665884. We thank Arnau Farr&#233; for critical reading of the manuscript; Maria Marsal for the help with the imaging and mounting of the 24 hpf embryo and; Senda Jim&#233;nez-Delgado for support with zebrafish micronjections.

The authors have nothing to disclose.

In this protocol, we describe a unique method to interrogate the mechanical properties of the cell nucleus inside the living cells. Different to other force spectroscopy techniques, non-invasive optical trapping allowed us to decouple the contribution of the cell membrane and cytoskeleton from the cell nuclear stiffness. Importantly, optical micromanipulation is compatible with multimodal microscopy, which will allow the experimenter to study different processes involved in cell nuclear mechanobiology. As a representative result, we used DNA-Hoechst staining to measure the nucleus deformation upon indentation performed by forces of the order of several hundreds of picoNewton.
Potential applications of our method beyond the examples outlined in this protocol 
The possibility to extract quantitative mechanical information from measurements inside living cells without external perturbations enable a plethora of unprecedented opportunities that are just beginning to be explored. Thus, the presented protocol of our optical micromanipulation platform can be extended to more complex experiments with great versatility. Acousto-optic deflectors (AOD) can generate multiple optical traps for synchronous force measurements across different cell locations, as well as can be used for active microrheology in a wide frequency range51,61. As has been mentioned, the force response upon indentation can overcome the maximum trapping force, leading to an escape of the bead from the optical trap. In this case, a force feedback can be configured with the AOD in order to clamp the optical force. All in all, multiple microrheological approaches, such as the stress relaxation described in this protocol, but also active microrheology or creep compliance, can be experimentally obtained with this platform and thoroughly analyzed by novel software packages61,62,63,64,65. Furthermore, the application of forces is not limited to the nucleus but could in principle be carried out to measure diverse intracellular structures and in complex tissues as demonstrated for trapping flowing red blood cells inside intact blood vessels66,67 or trapping and deforming chloroplasts and mitochondria68. Light-momentum calibration is independent of the shape and size of the trapped object, hence enabling direct force measurements on any force probe with arbitrary shape38,39. The use of injected microspheres allowed us to apply high forces onto the nucleus with relatively low laser power as compared to direct manipulation of cellular structures69,70,71. However, given a high enough refractive index difference, no externally applied force probe is necessary and intracellular organelles can be manipulated directly without injected beads (unpublished observations and reference70).
Potential modifications of our method to extend the applications 
Different sizes of microbeads can be injected depending on the experiment, but the relative controls must be done. For example, to study cells at later stages smaller beads can be injected. This will reduce the maximum force that can be exerted by the optical trap (such as shown in reference55). Bigger beads can be injected to exert higher forces, but these might affect embryo development depending on their size or stage of interest. In experiments where microbead injection is not an option, various organelles displaying refractive indices differences compared to that of the cytoplasm can still be optically manipulated, giving rise to optical forces measurable from light momentum changes42. As mentioned above, these methods have been employed by Bambardekar et al. to deform cell-cell junctions in the Drosophila embryo70. Likewise, the cell&#39;s nucleus has a lower refractive index than the surrounding medium44, which allows for bead-free indentation (unpublished observations and reference72) even though with a lower trapping strength. Thus, the nucleus cannot be trapped easily and escapes the trap.
The spin-coated PDMS spacer is fabricated via a convenient and fast method but might be out of reach for labs without access to a micro-/nanofabrication facility or engineering labs. Thus, the spacer can be easily assembled from lab tape or parafilm (step 4). The protocol can also be adapted by manufacturing microfluidic channels that automate the delivery of single cells into predefined measurement wells or into a chamber with a defined height to estimate the confinement effect within the same specimen. However, such microfluidic devices must be designed so that they fit the space between the microscope objective and the collecting lens of the optical force sensor, of around 2 mm (see step 3). Note that the optical force sensor must be positioned at the appropriate height so that no optical aberrations from defocusing affect the photon momentum measurement.
Other modifications could include the change of biological reporters. We found that Hoechst fluorescence spectrally bleeds into the GFP channel and we thus favor the combination with mCherry-tagged histone&#160;as a nuclear marker for simultaneous measurement in two fluorescent channels. Alternatively, nuclear deformation can easily be tracked with a label targeted to the inner nuclear membrane such as Lap2b-GFP (Figure 2).
Indentation onto the cell nucleus was of the order of 2-3 microns, which we could accurately measure by image analysis of diffraction-limited spinning-disk confocal microscopy. For the case of stiffer nuclei or smaller forces, indentation will be barely measurable using this approach. However, the&#160;absolute force-calibrated optical tweezers can be also calibrated for position measurements of the trapped bead in situ using BFP interferometry with nanometer accuracy51. Using this approach, the voltage signal and the optical force sensor can be translated into the position of the trapped probe through parameter&#160;&#946; [nm/V], while the invariant parameter&#160;&#945; [pN/V] yields force values through the aforementioned light-momentum calibration41 (see below for details).
Troubleshooting 
We found that the following challenges could occur during the experiment:
No stable trap is formed and the microsphere escapes easily 
Any dirt on the microscope objective or a misaligned correction collar could lead to a failure of a stable trap. If an immediate solution is not found, measure the point-spread function of the objective lens. If the specimen of interest is deep inside an optically dense tissue, the laser focus might experience severe optical aberrations leading to unstable trapping (this effect is usually negligible in isolated cells but becomes more evident in thicker tissues). For high stiffness, the restoring force of the nucleus could exceed the escape force of the trap, such that the microsphere is lost and the indentation routine fails. Initially, the nuclear membrane edge proximal to the optical trap gets hardly indented (Figure S2A). When this occurs, the trapping laser is no longer affected by force and Brownian motion, which leads to a force drop to zero and a decrease of the signal noise (Figure S2B). In case this happens, laser power can be increased to have a stronger trap, the amplitude of the trapezoidal trajectory pushing the bead into the nucleus can be reduced, or the initial position of the trapped microbead can be set further off the nucleus.
The cell is moving during the stimulation 
If cells are not sufficiently attached, the optical gradient trap will move the cells while performing the intracellular indentation routine, such that the forces and underlying mechanics of the nucleus are artefactual. To prevent displacement of the entire cell, we recommend increasing the concentration of cell adhesion molecules on the surface, e.g., ConA.
Initial momentum compensation 
If an initial momentum compensation routine is not available in the OTs platform (step 6.5), an artificial, force-independent baseline signal needs to be corrected for. This is visible as a slope on the force curve even with no bead trapped (Figure S1E). To do the correction, the same trajectory needs to be performed without a bead, outside of the cell at exactly the same position. For this, move the cell away from the trap using the stage control. As a reference, the force offset changes 5 pN across the FOV at 200 mW in our system; thus, it becomes negligible for short trajectories. Alternatively, a piezo scan stage can be used to move the cells on the sample, leaving the laser position constant.
Critical steps of the presented protocol 
Microspheres should be injected at the right, 1-cell stage to ensure maximal distribution over the embryo. Beads should not be fluorescent so that no light leaks into the fluorescent channels used for imaging. For example, even typical red-fluorescent beads are clearly visible in the blue channel used for imaging the cell nucleus after Hoechst staining due to their brightness (excitation: 405 nm; emission: 445 nm). Stable attachment of the cell to the substrate is critical to prevent lateral displacement during the indentation routine. If the cell moves during the routine, forces are underestimated. Should this happen frequently, optimize the attachment protocol. For tissue culture cells, other cell adhesion proteins, such as fibronectin, collagen, or poly-L-lysine lead to satisfactory attachment (unpublished observations). During the confinement, cells are subjected to a sudden and severe mechanical stress. This can cause damage to the cells and frequently the experimenter will encounter bursted cells if the procedure is not carried out carefully. Also, if the confinement height is too small, all the cells will suffer from nuclear envelop breakage or irreversible damage. To mitigate these, lower the upper coverslip more slowly and/or increase the spacing between the coverslip.
Limitations of the technique and suggestions to overcome them 
A clear limitation of the technique is the penetration of the laser light into deep sections of the tissue, which leads to aberrations and unstable trapping. Thus, a lower limit of penetration depth depends on the clarity of the sample, the aberration correction that can be employed73 and the applied laser power. It should be taken into account that a higher laser power leads to thermal excitation of the sample in the vicinity of the microsphere. However, heating of the sample originated by the 1064 nm wavelength laser spot is minimized to avoid plausible heat-related stress onto our biological samples74.
Another limitation is the maximum force that can be measured. Even though direct light-momentum detection enables force measurements far beyond the linear response regime of the optical trap40,41, the maximum applied force is in the order of a few hundred picoNewtons. This is limited by laser power and the consequential damage threshold of soft biological material and the refractive index differences, which are normally not larger than 0.1 or 0.344. Several methods have been proposed to increase the force detection limit, e.g., using structured light75, anti-reflective coated microspheres76, high-refractive index particles77 or highly doped quantum dots78.
OTs can be used for nanometer-scale position measurements through BFP interferometry, such that the position of the bead within the trap is&#160;&#916;x =&#160;&#160;&#946;Sx, where Sx is the voltage signal of the sensor, and&#160;&#946; [&#181;m/V] can be calibrated on-the-fly following different protocols35,54. For an optical force sensor, it can be proved that the voltage-to-force invariant conversion factor&#160;&#945; [pN/V] directly relates to&#160;&#946; and the trap stiffness, k [pN/&#181;m], through&#160;&#945; = k&#946; 37) In experiments with bead displacements that are too small to be detected from optical imaging, this strategy can be used to complement force measurements with small position detection. An example is the application of the experimental routines presented here onto very stiff nuclei, for which forces at reasonable laser powers (200-500 mW) are not sufficient to induce indentation values large enough. In that case, the bead needs to be brought in contact with the nucleus and the trapping stiffness must be calibrated prior to the measurement (step 8.6). The indentation d of the nucleus as a function of force can be indirectly determined as:
d = xtrap - F/k
where xtrap is the trap position. Different to the invariant light-momentum factor&#160;&#945; [pN/V], factor &#946; [&#181;m/V] needs to be calibrated prior to each experiment since it depends on many local variables determining the trapping dynamics, such as the particle size, optical trap spot size, and relative refractive indices.

Tissue morphogenesis is a complex process in which biochemical signals and physical forces are spatiotemporally coordinated. In the developing embryo, gradients of biochemical signaling factors dictate fate specification and ensure correct tissue patterning1,2. At the same time, intrinsic and extrinsic forces play a role in building the architecture of the embryo3,4. The influence of cell cortex mechanics in this context has been studied extensively5,6. The tight interconnection between mechano-chemical processes during morphogenesis relies on the properties of single cells to sense and respond to mechanical forces in their tissue microenvironment. Cells, thereby decode mechanical signals via the presence of force-sensitive subcellular and molecular elements that transduce mechanical information into specific signaling pathways controlling cell behavior, cell fate, and cell mechanics.
A hallmark of developmental processes is that cells organize as groups to build multicellular structures. As such, single cells rarely rearrange and move alone but are associated in a tight sociotope in which they show collective behavior such as supracellular migration7, (un)jamming transitions8,9 or blastocyst compaction10. Mechanical forces generated within and between cells serve as important cues to instruct collective cell dynamics7,11. But even when cells move alone, such as progenitor cells that squeeze their way between tissue sheets or narrow tissue niches, they experience extensive anisotropic mechanical forces when navigating a three-dimensional environment. These mechanical stresses on cells have profound consequences on cellular behavior12,13. Several mechanisms have been investigated that converge on the nucleus as a major mechanotransduction element14,15, as passive or active mechanical element during migration within a dense 3D tissue environment15,16.
We recently proposed a mechanism that equips cells to measure shape deformations using the nucleus as an elastic intracellular mechano-gauge12. The nucleus, being the largest organelle in a cell, undergoes large deformations when cells polarize, migrate, or change their shape under mechanical stretch, confinement, or osmotic stress16,17,18,19. We found that nuclear envelope stretch along with the intracellular positioning of the nucleus provides cells with information on the magnitude and type of cell deformation (such as cell compression versus cell swelling). Stretching of the nucleus is associated with an unfolding of the inner nuclear membrane (INM), which promotes calcium-dependent cPLA2 (cytosolic phospholipase A2) lipase activity at the INM followed by the release of Arachidonic Acid (AA) and rapid activation of myosin II at the cell cortex. This leads to increased cell contractility and amoeboid cell migration above a threshold of cortical contractility6. The mechanosensitive response to cell deformation occurs in less than a minute and is reversible upon confinement release, suggesting that the nucleus acts as a strain gauge for cellular proprioception regulating adaptive cell behavior under mechanical stress conditions. This mechanosensitive pathway is shown to be active in progenitor stem cells derived from zebrafish embryos, both in pluripotent and lineage-committed cells12 and is conserved in different species and cell lines20.
In addition to the nuclear properties as a cell-mechanosensor, nuclear architecture and mechanics are intrinsically regulated during development and in response to cell fate specification21, hence tuning cellular mechano-sensitivity22,23. The consequence might be a change in nuclear compliance that allows for morphological changes and transitions from a premigratory to a migratory state and vice versa8.
Several techniques to measure cell nucleus mechanics have been applied, such as atomic force microscopy24,25, micropipette aspiration26,27, microfluidic technology28, and microneedles29. However, many of these techniques are invasive in the sense that the entire cell must be deformed, limiting the measurement of mechanical characteristics and force-dependent responses of the nucleus itself. To circumvent the simultaneous deformation of the cell surface and its mechanosensitive cell cortex30, isolated nuclei were studied in various contexts31,32. However, it cannot be ruled out that nuclear isolation is associated with a change in mechanical nucleus properties and their regulation (reference24 and own unpublished observations).
Optical tweezers (OTs) are a versatile technology that has allowed a plethora of experiments in cell mechanobiology and have been instrumental in our understanding of how molecular machines convert chemical&#160;into mechanical energy33,34. Optical tweezers use a tightly focused laser beam to exert optical forces onto dielectric particles that have a refractive index higher than the surrounding medium33. Such forces can be of the order of hundreds of pico-Newtons and result in effective confinement of the particle within the laser trap focus, enabling manipulation of the trapped particle in three dimensions. The use of light has an important advantage in that the measurement can be performed non-invasively inside living cells. Optical manipulations are further limited to the trap focus of the laser beam. Hence, the manipulation can be performed without stimulating the surrounding cellular membranes and does not perturb the actin cortex or mechanosensitive processes at the plasma membrane, such as the force-dependent activation of ion channels.
The difficulty of the optical tweezer approach is to precisely determine the forces applied to the microsphere using classical approaches that rely on indirect force calibration based on the equipartition theorem or the use of defined Stokes-drag forces to measure a laser-power dependent escape force35. Whereas these methods are straightforward to implement in an in vitro experiment, they usually cannot be translated into a cellular environment. Several strategies have been introduced into the field that rely on a direct force calibration, derived from the first principles of momentum conservation36,37. Unlike other force spectroscopy approaches, force measurements are deduced from a local interchange of light momentum with the arbitrarily-shaped trapped particle38,39. In our experimental set-up, changes in light momentum arising from optical forces are directly measured without the need for in situ trap calibration40,41,42,43. Thus, the measurements become possible in a viscous environment such as the interior of the cell or even within a tissue, and forces can be readily quantified down to the pN level.
In this protocol, we describe an assay to mechanically manipulate intracellular organelles or structures and quantitatively assess their mechanical properties by an optical tweezer set-up. This set-up is integrated into a spinning disk fluorescent microscope enabling parallel imaging of cellular behavior or intracellular dynamics. The assay allows for the characterization of the mechanical properties of specific cellular compartments, such as the nucleus, while simultaneously studying the possible mechanoresponse and activation of molecular signaling pathways as a result of the deformation itself. Furthermore, optical trapping of injected microbeads within cells allows for an increase in the indentation force thanks to a considerably higher refractive index of the polystyrene bead (n = 1.59) compared to the intrinsic refractive contrast44 of the nucleus (n ~ 1.35) versus cytoplasm (n ~ 1.38). The presented strategy can be easily adapted to the study of other intracellular structures and organelles, as well as other approaches involving active microrheology, the use of multiple optical traps to probe the same/different sub-cellular structures simultaneously, and measurements targeting cell mechanobiology in the live embryo.

<table><tbody><tr><td>#1.5 22 mm cover glasses</td><td>Ted Pella</td><td>260148</td><td></td></tr><tr><td>#1.5 22x60 mm Coverglasses</td><td>Ted Pella</td><td>260152</td><td></td></tr><tr><td>#1.5H glass bottom dishes</td><td>Willco</td><td>GWST-5040</td><td></td></tr><tr><td>10-um beads</td><td>Supelco</td><td>72986</td><td></td></tr><tr><td>1-mm glass capillaries</td><td>Harvard Apparatus</td><td>30-0020 GC100F-15</td><td></td></tr><tr><td>1-um polystyrene microbeads</td><td>Sigma</td><td>89904</td><td></td></tr><tr><td>1-um red-fluorescent beads</td><td>ThermoFisher</td><td>F8816</td><td></td></tr><tr><td>Agar</td><td>ThermoFisher</td><td>16500500</td><td></td></tr><tr><td>Aqcuisition cameras sCMOS</td><td>Andor</td><td>Sona-4BV11-UNI</td><td></td></tr><tr><td>Auxiliary camera (Figure 3, AUX)</td><td>Blackfly, FLIR</td><td>BFS-U3-200S6M-C</td><td></td></tr><tr><td>Calbryte 520</td><td>AAT Bioquest</td><td>520 AM</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5453000011</td><td></td></tr><tr><td>Concanavalin A</td><td>Sigma</td><td>C5275</td><td></td></tr><tr><td>DMEM</td><td>Sigma</td><td>D2906</td><td></td></tr><tr><td>DNA-Hoechst</td><td>ThermoFisher</td><td>33342</td><td></td></tr><tr><td>Double scotch tape</td><td>Biesse Adesivi</td><td></td><td></td></tr><tr><td>E3</td><td>5 mM NaCl. 0.17 mM KCl. 0.33 mM CaCl2. 0.33 mM MgSO4</td><td></td><td></td></tr><tr><td>Eclipse Ti2</td><td>Nikon</td><td></td><td></td></tr><tr><td>Forceps</td><td>Fine Science Tools</td><td>11252-20</td><td></td></tr><tr><td>GPI-GFP</td><td></td><td></td><td></td></tr><tr><td>H2A-mCh</td><td></td><td></td><td></td></tr><tr><td>Image acquisition software</td><td>Fusion-Andor</td><td></td><td></td></tr><tr><td>Immersion Oil</td><td>Cargille</td><td>Type B: 16484</td><td></td></tr><tr><td>IR protection googles</td><td>Thorlabs</td><td>LG1</td><td></td></tr><tr><td>Lap2b-eGFP</td><td></td><td></td><td></td></tr><tr><td>Micro loader pipette</td><td>Eppendorf</td><td>GELoader</td><td></td></tr><tr><td>Microinjector</td><td>World Precision Instruments</td><td>SYS-PV820</td><td></td></tr><tr><td>MicroManager 2.0</td><td></td><td></td><td></td></tr><tr><td>Micromiter slide</td><td>ID5243 GXMGRAT-5 5mm/100 divisions</td><td></td><td></td></tr><tr><td>Mineral oil</td><td>Sigma</td><td>M3616</td><td></td></tr><tr><td>Motorized stage</td><td>ASI</td><td></td><td></td></tr><tr><td>Needle puller</td><td>Sutter instrument Co.</td><td>Model P-97</td><td></td></tr><tr><td>Optical tweezers platform</td><td>Impetux Optics</td><td>Sensocell</td><td></td></tr><tr><td>OTs software (LightAce)</td><td>Impetux Optics</td><td></td><td></td></tr><tr><td>PDMS</td><td>Sigma</td><td>Sylgard 184</td><td></td></tr><tr><td>PDMS Curing agent</td><td>Sigma</td><td>Sylgard 184</td><td></td></tr><tr><td>Post processing software (Matlab)</td><td>Mathworks</td><td></td><td></td></tr><tr><td>RNAse free water</td><td>Thermofisher</td><td>AM9937</td><td></td></tr><tr><td>Short-pass dichroic mirror (Figure 3, IR-F)</td><td>Semrock</td><td>FF01-950/SP-25</td><td></td></tr><tr><td>Spin-coater</td><td>Specialty Coating Systems</td><td>Spincoat G3P-8</td><td></td></tr><tr><td>Spinning-disk confocal microscope</td><td>Andor</td><td>DragonFly 502</td><td></td></tr><tr><td>Stereomicroscope</td><td>Leica M80</td><td></td><td></td></tr><tr><td>Triangular microinjection mold</td><td>Adaptive Science Tools</td><td>TU1</td><td></td></tr><tr><td>Universal oven</td><td>Memmert</td><td>UNB 200</td><td></td></tr><tr><td>Water immersion objective</td><td>Nikon</td><td>MRD07602</td><td></td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers at JoVE.com

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

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 ...

direct-force-measurements-subcellular-mechanics-confinement-using

Research

JoVE Journal

Developmental Biology

4.8K Views.  Institut de Ciències Fotòniques, ICFO. Dit protocol ontleedt de individuele mechanische eigenschappen van de celkern die het een mechanische graadmeter maken voor celpolarisatie, differentiatie en migratie. Het belangrijkste voordeel van deze techniek is dat het mechanische eigenschappen kan meten en krachten kan uitoefenen op subcellulaire compartimenten zonder externe verstoring van het membraan of corticale cytoskelet. Het protocol kan eenvoudig worden aangepast om kralen in andere cellen van andere organismen te manipuleren om...

Video: Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

Stretching Short Sequences of DNA with Constant Force Axial Optical Tweezers

Single-molecule techniques for stretching DNA of contour lengths less than a kilobase are fraught with experimental difficulties. However, many interesting biological events such as histone binding and protein-mediated looping of DNA1,2, occur on this length scale. In recent years, the mechanical properties of DNA have been shown to play a significant role in fundamental cellular processes like the packaging of DNA into compact nucleosomes and chromatin fibers3,4. Clearly, it is then important to understand the mechanical properties of short stretches of DNA. In this paper, we provide a practical guide to a single-molecule optical tweezing technique that we have developed to study the mechanical behavior of DNA with contour lengths as short as a few hundred basepairs.
The major hurdle in stretching short segments of DNA is that conventional optical tweezers are generally designed to apply force in a direction lateral to the stage5,6, (see Fig. 1). In this geometry, the angle between the bead and the coverslip, to which the DNA is tethered, becomes very steep for submicron length DNA. The axial position must now be accounted for, which can be a challenge, and, since the extension drags the microsphere closer to the coverslip, steric effects are enhanced. Furthermore, as a result of the asymmetry of the microspheres, lateral extensions will generate varying levels of torque due to rotation of the microsphere within the optical trap since the direction of the reactive force changes during the extension.
Alternate methods for stretching submicron DNA run up against their own unique hurdles. For instance, a dual-beam optical trap is limited to stretching DNA of around a wavelength, at which point interference effects between the two traps and from light scattering between the microspheres begin to pose a significant problem. Replacing one of the traps with a micropipette would most likely suffer from similar challenges. While one could directly use the axial potential to stretch the DNA, an active feedback scheme would be needed to apply a constant force and the bandwidth of this will be quite limited, especially at low forces.
We circumvent these fundamental problems by directly pulling the DNA away from the coverslip by using a constant force axial optical tweezers7,8. This is achieved by trapping the bead in a linear region of the optical potential, where the optical force is constant-the strength of which can be tuned by adjusting the laser power. Trapping within the linear region also serves as an all optical force-clamp on the DNA that extends for nearly 350 nm in the axial direction. We simultaneously compensate for thermal and mechanical drift by finely adjusting the position of the stage so that a reference microsphere stuck to the coverslip remains at the same position and focus, allowing for a virtually limitless observation period.

We illustrate the use of a constant force axial optical tweezers to explore the mechanical properties of short DNA molecules. By stretching DNA axially, we minimize steric hindrances and artifacts arising in conventional lateral manipulation, allowing us to study DNA molecules as short as ~100 nm.

Single-molecule techniques for stretching DNA of contour lengths  less than a kilobase are fraught with experimental difficulties. However, many ...

Bioengineering

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 ...

Micropipette Aspiration of Substrate-attached Cells to Estimate Cell Stiffness

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell proliferation, differentiation, migration and cell-cell interactions. The two key parameters of cellular biomechanics are cellular deformability or stiffness and the ability of the cells to contract and generate force. Here we describe a quick and simple method to estimate cell stiffness by measuring the degree of membrane deformation in response to negative pressure applied by a glass micropipette to the cell surface, a technique that is called Micropipette Aspiration or Microaspiration.
Microaspiration is performed by pulling a glass capillary to create a micropipette with a very small tip (2-50 &mu;m diameter depending on the size of a cell or a tissue sample), which is then connected to a pneumatic pressure transducer and brought to a close vicinity of a cell under a microscope. When the tip of the pipette touches a cell, a step of negative pressure is applied to the pipette by the pneumatic pressure transducer generating well-defined pressure on the cell membrane. In response to pressure, the membrane is aspirated into the pipette and progressive membrane deformation or "membrane projection" into the pipette is measured as a function of time. The basic principle of this experimental approach is that the degree of membrane deformation in response to a defined mechanical force is a function of membrane stiffness. The stiffer the membrane is, the slower the rate of membrane deformation and the shorter the steady-state aspiration length.The technique can be performed on isolated cells, both in suspension and substrate-attached, large organelles, and liposomes.
Analysis is performed by comparing maximal membrane deformations achieved under a given pressure for different cell populations or experimental conditions. A "stiffness coefficient" is estimated by plotting the aspirated length of membrane deformation as a function of the applied pressure. Furthermore, the data can be further analyzed to estimate the Young's modulus of the cells (E), the most common parameter to characterize stiffness of materials. It is important to note that plasma membranes of eukaryotic cells can be viewed as a bi-component system where membrane lipid bilayer is underlied by the sub-membrane cytoskeleton and that it is the cytoskeleton that constitutes the mechanical scaffold of the membrane and dominates the deformability of the cellular envelope. This approach, therefore, allows probing the biomechanical properties of the sub-membrane cytoskeleton.

Here we describe a quick and simple method to measure cell stiffness. The general principle of this approach is to measure membrane deformation in response to well-defined negative pressure applied through a micropipette to the cell surface. This method provides a powerful tool to study biomechanical properties of substrate-attached cells.

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell ...

Measurement of Tension Release During Laser Induced Axon Lesion to Evaluate Axonal Adhesion to the Substrate at Piconewton and Millisecond Resolution

The formation of functional connections in a developing neuronal network is influenced by extrinsic cues. The neurite growth of developing neurons is subject to chemical and mechanical signals, and the mechanisms by which it senses and responds to mechanical signals are poorly understood. Elucidating the role of forces in cell maturation will enable the design of scaffolds that can promote cell adhesion and cytoskeletal coupling to the substrate, and therefore improve the capacity of different neuronal types to regenerate after injury.
Here, we describe a method to apply simultaneous force spectroscopy measurements during laser induced cell lesion. We measure tension release in the partially lesioned axon by simultaneous interferometric tracking of an optically trapped probe adhered to the membrane of the axon. Our experimental protocol detects the tension release with piconewton sensitivity, and the dynamic of the tension release at millisecond time resolution. Therefore, it offers a high-resolution method to study how the mechanical coupling between cells and substrates can be modulated by pharmacological treatment and/or by distinct mechanical properties of the substrate.

We measured the tension release in an axon that was partially lesioned with a laser dissector by simultaneous force spectroscopy measurement performed on an optically-trapped probe adhered to the membrane of the axon. The developed experimental protocol evaluates the axon adhesion to the culture substrate.

The formation of functional connections in a developing neuronal network is influenced by extrinsic cues. The neurite growth of developing neurons is ...

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to understand morphogenetic processes is to directly measure cellular forces and mechanical properties in vivo during embryogenesis. Here, we present a setup of optical tweezers coupled to a light sheet microscope, which allows to directly apply forces on cell-cell contacts of the early Drosophila embryo, while imaging at a speed of several frames per second. This technique has the advantage that it does not require the injection of beads into the embryo, usually used as intermediate probes on which optical forces are exerted. We detail step by step the implementation of the setup, and propose tools to extract mechanical information from the experiments. By monitoring the displacements of cell-cell contacts in real time, one can perform tension measurements and investigate cell contacts&#39; rheology.

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

We present a setup of optical tweezers coupled to a light sheet microscope, and its implementation to probe cell mechanics without beads in the Drosophila embryo.

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to ...

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the averaging that takes place in bulk-phase studies. To achieve visualization, dual optical tweezers, where one trap is fixed and the other is mobile, are focused into one channel of a multi-stream microfluidic chamber positioned on the stage of an inverted fluorescence microscope. The optical tweezers trap single molecules of fluorescently labeled DNA and fluid flow through the chamber and past the trapped beads, stretches the DNA to B-form (under minimal force, i.e., 0 pN) with the nucleic acid being observed as a white string against a black background. DNA molecules are moved from one stream to the next, by translating the stage perpendicular to the flow to enable the initiation of reactions in a controlled manner. To achieve success, microfluidic devices with optically clear channels are mated to glass syringes held in place in a syringe pump. Optimal results use connectors permanently bonded to the flow cell, tubing that is mechanically rigid and chemically resistant, and which is connected to switching valves that eliminate bubbles that prohibit laminar flow.

Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the ...

Biochemistry

High-Speed Magnetic Tweezers for Nanomechanical Measurements on Force-Sensitive Elements

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are therefore poised to be useful in the field of mechanobiology. Since the method commonly relies on image-based tracking of magnetic beads, the speed limit in recording and analyzing images, as well as the thermal fluctuations of the beads, has long hampered its application in observing small and fast structural changes in target molecules. This article describes detailed methods for the construction and operation of a high-resolution MT setup that can resolve nanoscale, millisecond dynamics of biomolecules and their complexes. As application examples, experiments with DNA hairpins and SNARE complexes (membrane-fusion machinery) are demonstrated, focusing on how their transient states and transitions can be detected in the presence of piconewton-scale forces. We expect that high-speed MTs will continue to enable high-precision nanomechanical measurements on molecules that sense, transmit, and generate forces in cells, and thereby deepen our molecular-level understanding of mechanobiology.

Here, we describe a high-speed magnetic tweezer setup that performs nanomechanical measurements on force-sensitive biomolecules at the maximum rate of 1.2 kHz. We introduce its application to DNA hairpins and SNARE complexes as model systems, but it will be also applicable to other molecules involved in mechanobiological events.

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are ...

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 ...

Probing Myosin Ensemble Mechanics in Actin Filament Bundles Using Optical Tweezers

Myosins are motor proteins that hydrolyze ATP to step along actin filament (AF) tracks and are essential in cellular processes such as motility and muscle contraction. To understand their force-generating mechanisms, myosin II has been investigated both at the single-molecule (SM) level and as teams of motors in vitro using biophysical methods such as optical trapping.
These studies showed that myosin force-generating behavior can differ greatly when moving from the single-molecule level in a three-bead arrangement to groups of motors working together on a rigid bead or coverslip surface in a gliding arrangement. However, these assay constructions do not permit evaluating the group dynamics of myosin within viscoelastic structural hierarchy as they would within a cell. We have developed a method using optical tweezers to investigate the mechanics of force generation by myosin ensembles interacting with multiple actin filaments.
These actomyosin bundles facilitate investigation in a hierarchical and compliant environment that captures motor communication and ensemble force output. The customizable nature of the assay allows for altering experimental conditions to understand how modifications to the myosin ensemble, actin filament bundle, or the surrounding environment result in differing force outputs.

Formation of actomyosin bundles in vitro and measuring myosin ensemble force generation using optical tweezers is presented and discussed.

Myosins are motor proteins that hydrolyze ATP to step along actin filament (AF) tracks and are essential in cellular processes such as motility and muscle ...

Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultrafast Force-Clamp Spectroscopy

Ultrafast force-clamp spectroscopy (UFFCS) is a single molecule technique based on laser tweezers that allows the investigation of the chemomechanics of both conventional and unconventional myosins under load with unprecedented time resolution. In particular, the possibility to probe myosin motors under constant force right after the actin-myosin bond formation, together with the high rate of the force feedback (200 kHz), has shown UFFCS to be a valuable tool to study the load dependence of fast dynamics such as the myosin working stroke. Moreover, UFFCS enables the study of how processive and non-processive myosin-actin interactions are influenced by the intensity and direction of the applied force.
By following this protocol, it will be possible to perform ultrafast force-clamp experiments on processive myosin-5 motors and on a variety of unconventional myosins. By some adjustments, the protocol could also be easily extended to the study of other classes of processive motors such as kinesins and dyneins. The protocol includes all the necessary steps, from the setup of the experimental apparatus to sample preparation, calibration procedures, data acquisition and analysis.

Presented here is a comprehensive protocol to perform ultrafast force-clamp experiments on processive myosin-5 motors, which could be easily extended to the study of other classes of processive motors. The protocol details all the necessary steps, from the setup of the experimental apparatus to sample preparation, data acquisition and analysis.

Ultrafast force-clamp spectroscopy (UFFCS) is a single molecule technique based on laser tweezers that allows the investigation of the chemomechanics of ...

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

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