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.

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			<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>
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		<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>
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		<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>
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		<tr>
			<td>1-um red-fluorescent beads</td>
			<td>ThermoFisher</td>
			<td>F8816</td>
			<td></td>
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		<tr>
			<td>Agar</td>
			<td>ThermoFisher</td>
			<td>16500500</td>
			<td></td>
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		<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>
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		<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>
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		<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>
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		<tr>
			<td>Eclipse Ti2</td>
			<td>Nikon</td>
			<td></td>
			<td></td>
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		<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>
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		<tr>
			<td>H2A-mCh</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr>
			<td>Image acquisition software</td>
			<td>Fusion-Andor</td>
			<td></td>
			<td></td>
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		<tr>
			<td>Immersion Oil</td>
			<td>Cargille</td>
			<td>Type B: 16484</td>
			<td></td>
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		<tr>
			<td>IR protection googles</td>
			<td>Thorlabs</td>
			<td>LG1</td>
			<td></td>
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		<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>
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		<tr>
			<td>Mineral oil</td>
			<td>Sigma</td>
			<td>M3616</td>
			<td></td>
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		<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>
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		<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>
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		<tr>
			<td>Stereomicroscope</td>
			<td>Leica M80</td>
			<td></td>
			<td></td>
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		<tr>
			<td>Triangular microinjection mold</td>
			<td>Adaptive Science Tools</td>
			<td>TU1</td>
			<td></td>
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		<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>
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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

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JoVE Journal

Developmental Biology

4.6K Views.  Institut de Ciències Fotòniques, ICFO. 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.

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

Direct Induction of Hemogenic Endothelium and Blood by Overexpression of Transcription Factors in Human Pluripotent Stem Cells

During development, hematopoietic cells arise from a specialized subset of endothelial cells, hemogenic endothelium (HE). Modeling HE development in vitro is essential for mechanistic studies of the endothelial-hematopoietic transition and hematopoietic specification. Here, we describe a method for the efficient induction of HE from human pluripotent stem cells (hPSCs) by way of overexpression of different sets of transcription factors. The combination of ETV2 and GATA1 or GATA2 TFs is used to induce HE with pan-myeloid potential, while a combination of GATA2 and TAL1 transcription factors allows for the production of HE with erythroid and megakaryocytic potential. The addition of LMO2 to GATA2 and TAL1 combination substantially accelerates differentiation and increases erythroid and megakaryocytic cells production. This method provides an efficient and rapid means of HE induction from hPSCs and allows for the observation of the endothelial-hematopoietic transition in a culture dish. The protocol includes hPSCs transduction procedures and post-transduction analysis of HE and blood progenitors.

This protocol describes the efficient induction of hemogenic endothelium and multipotential hematopoietic progenitors from human pluripotent stem cells via the forced expression of transcription factors.

During development, hematopoietic cells arise from a specialized subset of endothelial cells, hemogenic endothelium (HE). Modeling HE development in vitro ...

Using Isolated Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High throughput Microplate Respiratory Measurements

Skeletal muscle mitochondria play a specific role in many disease pathologies. As such, the measurement of oxygen consumption as an indicator of mitochondrial function in this tissue has become more prevalent. Although many technologies and assays exist that measure mitochondrial respiratory pathways in a variety of cells, tissue and species, there is currently a void in the literature in regards to the compilation of these assays using isolated mitochondria from mouse skeletal muscle for use in microplate based technologies. Importantly, the use of microplate based respirometric assays is growing among mitochondrial biologists as it allows for high throughput measurements using minimal quantities of isolated mitochondria. Therefore, a collection of microplate based respirometric assays were developed that are able to assess mechanistic changes/adaptations in oxygen consumption in a commonly used animal model. The methods presented herein provide step-by-step instructions to perform these assays with an optimal amount of mitochondrial protein and reagents, and high precision as evidenced by the minimal variance across the dynamic range of each assay.

The methods presented provide step-by-step instructions for the performance of a collection of microplate based respirometric assays using isolated mitochondria from minimal quantities of mouse skeletal muscle. These assays are able to measure mechanistic changes/adaptations in mitochondrial oxygen consumption in a commonly used animal model.

Skeletal muscle mitochondria play a specific role in many disease pathologies. As such, the measurement of oxygen consumption as an indicator of ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Protocol for the Direct Conversion of Murine Embryonic Fibroblasts into Trophoblast Stem Cells

Trophoblast stem cells (TSCs) arise as a consequence of the first cell fate decision in mammalian development. They can be cultured in vitro, retaining the ability to self-renew and to differentiate into all subtypes of the trophoblast lineage, equivalent to the in vivo stem cell population giving rise to the fetal portion of the placenta. Therefore, TSCs offer a unique model to study placental development and embryonic versus extra-embryonic cell fate decision in vitro. From the blastocyst stage onwards, a distinct epigenetic barrier consisting of DNA methylation and histone modifications tightly separates both lineages. Here, we describe a protocol to fully overcome this lineage barrier by transient over-expression of trophoblast key regulators Tfap2c, Gata3, Eomes and Ets2 in murine embryonic fibroblasts. The induced trophoblast stem cells are able to self-renew and are almost identical to blastocyst derived trophoblast stem cells in terms of morphology, marker gene expression and methylation pattern. Functional in vitro and in vivo assays confirm that these cells are able to differentiate along the trophoblast lineage generating polyploid trophoblast giant cells and chimerizing the placenta when injected into blastocysts. The induction of trophoblast stem cells from somatic tissue opens new avenues to study genetic and epigenetic characteristics of this extra-embryonic lineage and offers the possibility to generate trophoblast stem cell lines without destroying the respective embryo.

Here we present a protocol for the direct conversion of murine embryonic fibroblasts into fully functional and stable trophoblast stem cells by ten day over-expression of Tfap2c, Gata3, Eomes and Ets2.

Trophoblast stem cells (TSCs) arise as a consequence of the first cell fate decision in mammalian development. They can be cultured in vitro, retaining ...

Subcutaneous Neurotrophin 4 Infusion Using Osmotic Pumps or Direct Muscular Injection Enhances Aging Rat Laryngeal Muscles

Laryngeal dysfunction in the elderly is a major cause of disability, from voice disorders to dysphagia and loss of airway protective reflexes. Few, if any, therapies exist that target age-related laryngeal muscle dysfunction. Neurotrophins are involved in muscle innervation and differentiation of neuromuscular junctions (NMJs). It is thought that neurotrophins enhance neuromuscular transmission by increasing neurotransmitter release. The neuromuscular junctions (NMJs) become smaller and less abundant in aging rat laryngeal muscles, with evidence of functional denervation. We explored the effects of NTF4 for future clinical use as a therapeutic to improve function in aging human laryngeal muscles. Here, we provide the detailed protocol for systemic application and direct injection of NTF4 to investigate the ability of aging rat laryngeal muscle to remodel in response to NTF4 application. In this method, rats either received NTF4 either systemically via osmotic pump or by direct injection through the vocal folds. Laryngeal muscles were then dissected and used for histological examination of morphology and age-related denervation.

Here, we present a protocol to describe the use of neurotrophin 4 (NTF4) systemically and directly to remodel rat aging laryngeal muscles.

Laryngeal dysfunction in the elderly is a major cause of disability, from voice disorders to dysphagia and loss of airway protective reflexes. Few, if ...

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 ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

Direct Lineage Reprogramming of Adult Mouse Fibroblast to Erythroid Progenitors

Erythroid cell commitment and differentiation proceed through activation of a lineage-restricted transcriptional network orchestrated by a group of cell fate determining and maturing factors. We previously set out to define the minimal set of factors necessary for instructing red blood cell development using direct lineage reprogramming of fibroblasts into induced erythroid progenitors/precursors (iEPs). We showed that overexpression of Gata1, Tal1, Lmo2, and c-Myc (GTLM) can rapidly convert murine and human fibroblasts directly to iEPs that resemble bona fide erythroid cells in terms of morphology, phenotype, and gene expression. We intend that iEPs will provide an invaluable tool to study erythropoiesis and cell fate regulation. Here we describe the stepwise process of converting murine tail tip fibroblasts into iEPs via transcription factor-driven direct lineage reprogramming (DLR). In this example, we perform the reprogramming in fibroblasts from erythroid lineage-tracing mice that express the yellow fluorescent protein (YFP) under the control of the erythropoietin receptor gene (EpoR) promoter, enabling visualization of erythroid cell fate induction upon reprogramming. Following this protocol, fibroblasts can be reprogrammed into iEPs within five to eight days. 
While improvements can still be made to the process, we show that GTLM-mediated reprogramming is a rapid and direct process, yielding cells with properties of bona fide erythroid progenitor and precursor cells.

Here we present our protocol for producing induced erythroid progenitors (iEPs) from mouse adult fibroblasts using transcription factor-driven direct lineage reprogramming (DLR).

Erythroid cell commitment and differentiation proceed through activation of a lineage-restricted transcriptional network orchestrated by a group of cell ...

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...