This work was supported by an FRM Equipe Grant FRM DEQ20130326509, Agence Nationale de la Recherche ANR-Blanc Grant, Morfor ANR-11-BSV5-0008 (to P.-F.L.). We acknowledge France-BioImaging infrastructure supported by the French National Research Agency (ANR-10-INBS-04-01, &#171;Investments for the future&#187;). We thank Brice Detailleur and Claude Moretti from the PICSL-FBI infrastructure for technical assistance.

<ol>
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The authors have nothing to disclose.

Optical tweezers allow to perform absolute mechanical measurements directly in the developing embryonic epithelium in a non-invasive manner. In that sense, it presents advantages over other methods such as laser ablation, which are invasive and provide relative measurements, magnetic forces, which require injections, or force inference, which relies on strong assumptions and also provide relative measurements.
The protocol includes a few critical steps. First, as the objective lens may show chromatic aberrations and that the laser trap pushes the object &#34;downstream&#34;, it is important to check that the IR laser traps the bead in the imaging plane, and eventually correct for it (step 2.18). Second, the method relies on the measurements of the cell contacts&#39; position. It is thus crucial to use a high-contrast fluorescent marker.
The quality of the objective lens and the laser beam are critical to effective trapping. The numerical aperture of the objective lens should exceed 1.0. If trapping is ineffective, make sure that the laser beam fills the rear aperture of the objective lens.
Our method comes with several limitations. First, it is not clear what exactly provides the support for optical trapping. Although a mismatch of refractive index is detected, its origin remains to be determined. Second, the calibration process, if one is interested in absolute measurements, can be a bit tedious, as it requires passive microrheology experiments, and calibration of the trap stiffness on beads. It is important to recognize that the calibration of the stiffness on cell contacts is subject to experimental uncertainty: it relies on measurements of the trap stiffness on beads, which can be measured only in the cytosol, but not near cell contacts. Third, it is unclear how versatile optical tweezers can be. Although they are able to deform the cells in the Drosophila embryo, other tissues might present higher tension or more difficult access (such as going through a cuticle) and therefore be less amenable to optical tweezing.
Optical forces are small (&#60;few tens of pN) and may be thus insufficient to deform stiff or highly tensed structures. Magnetic tweezers on large particles would probably be more effective in this case.
We described here the coupling of optical tweezers to a light sheet microscope, but optical tweezers can be coupled to other types of microscopes, such as an epifluorescence or a confocal spinning disk microscope. The introduction of the IR trapping laser into the microscope depends on the microscope configuration. It mainly requires the possibility to add a dichroic mirror to combine the paths for imaging and optical manipulation. Most microscopy companies propose modular illumination systems, with a two-layer module, which allow this combination.
Several directions exist to improve or upgrade the technique. A possibility is to split the laser residence time among several positions or to use more advanced holographic techniques, to produce several traps. This could allow to create more complex force patterns on target cells or cell contacts. Another improvement could be to design a real-time feedback between the deflection caused and the position of the trap. This could allow proper creep experiments in which the force applied is maintained constant throughout the experiment.

Embryonic development is a highly reproducible process during which the cells and tissues deform to shape the future animal. Such deformations have been shown to require the active generation of forces at the cell level1,2. To understand morphogenetic processes during which cells and tissues change their shape, it is therefore key to assess the mechanical properties of the cells involved, and to measure the forces within the tissue during the process3,4. Epithelial layers, especially in Drosophila, have been widely studied due to their quasi-2D geometry and to their easy manipulation.
A number of techniques have thus been developed by us and others to assess epithelial mechanics in vivo during development. We will give a quick overview of the three main techniques used in epithelial tissues. Laser ablation, a widely used method, allows to reveal the local mechanical stress at cell junctions5,6,7,8 or at larger scale9,10,11 by performing local cuts that disrupt the mechanical integrity of the target. The dynamics of the opening following the cut provides information both on the stress prior ablation, and on the rheology of the tissue12,13. A drawback of laser ablation is that it is invasive, as it requires the local disruption of the cell cortex. Hence, one can only perform a limited number of ablations if one wants to preserve tissue integrity. Another drawback is that ablations only provide relative estimates of tension at cell contacts, since the opening velocity is dependent on viscous friction, which is generally not known. Magnetic manipulation has also been developed and used in Drosophila, involving either the use of ferrofluids14 or ultramagnetic liposomes15. They can provide absolute measurements16,17, but are also invasive in the sense that they require the injection of probes at the desired location. This can be very tricky depending on the system, which is not always amenable to precise injections. A third technique, fully non-invasive, is force inference18,19,20. Force inference relies on the assumption of mechanical equilibrium at triple points (tricellular junctions, or vertices), and allows to infer tensions at all cell-cell contacts (and possibly, pressures in all cells) by solving an inverse problem. For tensions, each vertex provides two equations (X and Y). This yields a large system of linear equations which can be inverted under some conditions to assess tension at all cell contacts. While this method is very attractive, as it only requires a segmented image and no extra experiment or setup, its accuracy is yet to determine, and again it only provides relative values, unless an absolute calibration measurement is performed.
To overcome some of these limitations, we introduce in this article a setup of optical tweezers coupled to a light sheet microscope to apply controlled forces at the cell scale in the embryonic epithelium of Drosophila melanogaster. Optical tweezers have been used for numerous biological applications including the measurements on single proteins and manipulation of organelles and cells21. Here, we report applied forces in the range of a few dozen pN, which is small yet sufficient to induce local deformations of cell contacts and perform mechanical measurements in vivo. Typically, we use perpendicular deflection of cell contacts, monitored through the analysis of kymographs, to relate force to deformation. Importantly, our setup does not require the injection of beads at the desired location in the tissue, as optical tweezers are able to directly exert forces on cell-cell contacts. The coupling of the optical tweezers to a light sheet microscope allows one to perform rapid imaging (several frames per second), which is very appreciable for a mechanical analysis at short time scales, and with reduced phototoxicity, since the illumination of the sample is limited to the plane of imaging22.
Overall, optical tweezers are a non-invasive way to apply controlled forces at cell contacts in vivo in the Drosophila embryo, and to extract mechanical information such as stiffness and tension at cell contacts23, rheological properties24, and gradient or anisotropy of tension23.

<table><tbody><tr><td>Ytterbium Fiber Laser LP, 10 W, CW</td><td>IPG Laser</td><td>YLM-10-LP-SC</td><td>including collimator LP: beam D = 1.6 mm and red guide laser</td></tr><tr><td>&amp;Oslash;1/2&quot; Optical Beam Shutter</td><td>Thorlabs</td><td>SH05</td><td></td></tr><tr><td>Small Beam Diameter Galvanometer Systems</td><td>Thorlabs</td><td>GVS001</td><td>1 for X displacement, 1 for Y displacement</td></tr><tr><td>1D or 2D Galvo System Linear Power Supply</td><td>Thorlabs</td><td>GPS011</td><td>galvanometers power supply</td></tr><tr><td>2 lenses f = 30 mm</td><td>Thorlabs</td><td>LB1757-B</td><td>relay telescope between 2 galva</td></tr><tr><td>Lens f = 200 mm</td><td>Thorlabs</td><td>LB1945-B</td><td>2.5X telescope</td></tr><tr><td>Lens f = 500 mm</td><td>Thorlabs</td><td>LB1869-B</td><td>2.5X telescope</td></tr><tr><td>Right-Angle Kinematic Elliptical Mirror Mount with Tapped Cage Rod Holes</td><td>Thorlabs</td><td>KCB1E</td><td>Periscope</td></tr><tr><td>Laser Safety Glasses, Light Green Lenses, 59% Visible Light Transmission, Universal Style</td><td>Thorlabs</td><td>LG1</td><td></td></tr><tr><td>45&amp;deg; AOI, 50.0 mm Diameter, Hot Mirror</td><td>Edmund Optics</td><td>#64-470</td><td></td></tr><tr><td>Multiphoton-Emitter HC 750/S</td><td>AHF</td><td>HC 750/SP</td><td></td></tr><tr><td>CompactDAQ Chassis</td><td>National Instruments</td><td>cDAQ-9178</td><td></td></tr><tr><td>C Series Voltage Output Module</td><td>National Instruments</td><td>NI-9263</td><td>Analog output module</td></tr><tr><td>C Series Voltage Input Module</td><td>National Instruments</td><td>NI-9215</td><td>Analog input module</td></tr><tr><td>FluoSpheres Carboxylate-Modified Microspheres, 0.5 &amp;micro;m, red fluorescent (580/605), 2% solids</td><td>ThermoFisher Scientific</td><td>F8812</td><td>calibration beads</td></tr><tr><td>C++ (Qt) home made optical tweezers software</td><td></td><td></td><td>developed by Olivier Blanc and Claire Chard&amp;egrave;s. Alternative solution: labview</td></tr></tbody></table>

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.

Figure 5 shows experimental data obtained by imposing a sinusoidal movement to the trap. The trap produces a deflection of the interface, as exemplified by the 3 snapshots showing 3 successive interface positions (Figure 5A)23. Recorded movies are used to generate a kymograph (Figure 5B) and are further analyzed to determine the position of the interface with subpixel resolution, using a Gaussian fit along the x direction for each frame. In the regime of small deformations, the trap and interface positions are proportional (Figure 5C). The amplitude of the interface deflection relative to that of the trap (Figure 5D) gives access to the interface tension or stiffness (see step 7.1).
Figure 6 shows experimental data obtained by performing pull release experiments. The optical trap is switched on approximately 1 &#181;m away from the midpoint of the interface between two cells, which causes the interface to deflect towards the trap position (Figure 6A)24. The trap is then switched off after the desired amount of time. The position xm of the interface (Figure 6B) is obtained from kymographs (Figure 6C), again using Gaussian fits along the x direction for each frame. The resulting graph can be compared to hypothesized rheological models, for instance, a Maxwell model (Figure 6D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57900/57900fig1.jpg" /> 
Figure 1: Schematic of the optical tweezers (red path) setup combined with the light sheet microscope. This figure has been modified from Bambardekar, K. et al.23.The light sheet microscope, composed by the illumination unit and the detection unit, is described previously25. The optical tweezers correspond to the red part of the scheme: The infrared laser passes through an optical shutter and 2 galvanometers which control the position of the trap in the sample. A 1-fold telescope is placed between the 2 galvanometers to keep the conjugation between them. Then, a telescope increases the size of the beam by 2.5-fold and the periscope brings it to the height of the microscope. The infrared laser enters the detection objective of the microscope thanks to a dichroic mirror.Important distances between optical elements are given directly in the figure. Distance between the last lens (focal length 500 mm) and the back aperture of the objective is 500 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57900/57900fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57900/57900fig2.jpg" /> 
Figure 2: Homemade modified rail holding the dichroic mirror and the hot mirror. The dichroic mirror rail of the microscope has been cut on the left side to allow the entrance of the infrared laser. The dichroic mirror reflects the infrared light and transmits the visible light (fluorescence). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57900/57900fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57900/57900fig3.jpg" /> 
Figure 3: Connection of the instruments to the NI Cards. AO: analog output, AI: analog input, AO0 and AI0 are connected to galvo1, AO1 and AI1 are connected to galvo2, PFI0 is connected to the fire of the camera, to AI2 and PFI1 is connected to the trigger in of the shutter and AI3 is connected to the trigger out of the shutter. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57900/57900fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57900/57900fig4.jpg" /> 
Figure 4: Sample holder in the observation context. This figure has been modified from Chard&#232;s, C., et al.25. The embryos are immobilized on the glass coverslip. The slide is held by the sample holder. The sample holder is inserted in the glass cuvette, held by the holder fixed to the piezoelectric stage. The light sheet is horizontal and illuminates the embryos from the side. The detection objective lens is vertical, above the sample, and dips into the cuvette.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57900/57900fig5.jpg" /> 
Figure 5: Interface deflection imposed by sinusoidal movement of the trap. This figure has been modified from Bambardekar, K. et al23. (A) The trap is moved sinusoidally perpendicular to the interface. The trap and interface positions are xt and xm, respectively. The right panels show three images of the interface at different positions. The laser trap position is marked by a yellow arrowhead. Interface are labelled with a membrane marker (GAP43::mcherry). (B) Kymograph along the axis defined by the direction of trap movement (perpendicular to the interface) (Period = 5 s). (C) Representative plot of deflection versus time showing both trap (red solid line) and interface positions (black solid line). (D) Interface position as a function of trap position during few cycles of laser oscillation (Amplitude = 0.5 &#181;m, Period = 2 s). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57900/57900fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/57900/57900fig6.jpg" /> 
Figure 6: Interface deflection in pull-release experiments. This figure has been modified from Serge, A., et al.24. (A) The trap is switched on at a distance from the midpoint of the interface, then switched off again. (B) The trap and interface positions are xt and xm, respectively. Kymographs are generated along the x direction, perpendicular to the contact line&#39;s midpoint. (C) Kymograph of a pull-release experiment. Cell contacts are labelled with Utrophin::GFP. (D) Representative plot of deflection versus time showing both trap (dotted red/green lines) and interface position (black solid line). The solid red line shows a fit obtained using a Maxwell-like rheological model24. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57900/57900fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Movie 1: Tweezing experiment. Pixel size = 194 nm, 10 fps, Trap oscillation period = 2 s, Labelling: Gap43::mCherry. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57900/Movie1_crop-Gap43MC-10fps-2sOsc.tif">Please click here to download this file.</a>

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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.

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 ...

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Nanyang Technological University

Research

JoVE Journal

Developmental Biology

7.5K Views.  Aix-Marseille Université. Deze methode kan helpen bij het beantwoorden van belangrijke vragen in weefselmechanica, zoals hoe eigenschappen van celcelcontacten veranderen in ruimte en tijd tijdens weefselmorfogenese. Het belangrijkste voordeel van deze techniek is dat het zwak invasief is en compatibel is met live beeldvorming zoals lichtplaatmicroscopie. Deze techniek vereist geen injectie van kralen in het weefsel.Meestal gebruikt als demiere sondes waarop froses te krijgen, ik ben opgewonden. Het aantonen van...

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

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

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

Neuroscience

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila larvae, customized metal nickel-titanium alloy (nitinol) filaments. These mechanical probes are similar to the von Frey filaments used in vertebrates to measure mechanical nociception. Here, we demonstrate how to make and calibrate these mechanical probes and how to generate a full behavioral dose-response from subthreshold (innocuous or non-noxious range) to suprathreshold (low to high noxious range) stimuli. To demonstrate the utility of the probes, we investigated tissue damage-induced hypersensitivity in Drosophila larvae. Mechanical allodynia (hypersensitivity to a normally innocuous mechanical stimulus) and hyperalgesia (exaggerated responsiveness to a noxious mechanical stimulus) have not yet been established in Drosophila larvae. Using mechanical probes that are normally innocuous or probes that typically elicit an aversive behavior, we found that Drosophila larvae develop mechanical hypersensitization (both allodynia and hyperalgesia) after tissue damage. Thus, the mechanical probes and assay that we illustrate here will likely be important tools to dissect the fundamental molecular/genetic mechanisms of mechanical hypersensitivity.

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

The goal of this protocol is to show how to perform an improved assay for mechanical nociception in Drosophila larvae. We use the assay here to demonstrate that mechanical hypersensitivity (allodynia and hyperalgesia) exists in Drosophila larvae.

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila ...

Behavior

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

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

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

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

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

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

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 ...

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of embryogenesis, these organelles undergo dramatic movements powered by actin-based cytoplasmic streaming and motor-driven trafficking along microtubules. The development of a multitude of small, organelle-specific fluorescent probes (FPs) makes it possible to visualize a wide range of different lipid-containing structures in any genotype, allowing live imaging without requiring a genetically encoded fluorophore. This protocol shows how to inject vital dyes and molecular probes into Drosophila embryos to monitor the trafficking of specific organelles by live imaging. This approach is demonstrated by labeling lipid droplets (LDs) and following their bulk movement by particle image velocimetry (PIV). This protocol provides a strategy amenable to the study of other organelles, including lysosomes, mitochondria, yolk vesicles, and the ER, and for tracking the motion of individual LDs along microtubules. Using commercially available dyes brings the benefits of separation into the violet/blue and far-red regions of the spectrum. By multiplex co-labeling of organelles and/or cytoskeletal elements via microinjection, all the genetic resources in Drosophila are available for trafficking studies without the need to introduce fluorescently tagged proteins. Unlike genetically encoded fluorophores, which have low quantum yields and bleach easily, many of the available dyes allow for rapid and simultaneous capture of several channels with high photon yields.

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of ...

Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility in Drosophila

Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility Coupled with Measurement of Epithelial Tension in Drosophila Embryos

Contractile forces generated by actin and non-muscle myosin II ("actomyosin contractility") are critical for morphological changes of cells and tissues at multiple length scales, such as cell division, cell migration, epithelial folding, and branching morphogenesis. An in-depth understanding of the role of actomyosin contractility in morphogenesis requires approaches that allow the rapid inactivation of actomyosin, which is difficult to achieve using conventional genetic or pharmacological approaches. The presented protocol demonstrates the use of a CRY2-CIBN based optogenetic dimerization system, Opto-Rho1DN, to inhibit actomyosin contractility in Drosophila embryos with precise temporal and spatial controls. In this system, CRY2 is fused to the dominant negative form of Rho1 (Rho1DN), whereas CIBN is anchored to the plasma membrane. Blue light-mediated dimerization of CRY2 and CIBN results in rapid translocation of Rho1DN from the cytoplasm to the plasma membrane, where it inactivates actomyosin by inhibiting endogenous Rho1. In addition, this article presents a detailed protocol for coupling Opto-Rho1DN-mediated inactivation of actomyosin with laser ablation to investigate the role of actomyosin in generating epithelial tension during Drosophila ventral furrow formation. This protocol can be applied to many other morphological processes that involve actomyosin contractility in Drosophila embryos with minimal modifications. Overall, this optogenetic tool is a powerful approach to dissect the function of actomyosin contractility in controlling tissue mechanics during dynamic tissue remodeling.

Author Spotlight: Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility Coupled with Measurement of Epithelial Tension in Drosophila Embryos  

Actomyosin contractility plays an important role in cell and tissue morphogenesis. However, it is challenging to manipulate actomyosin contractility in vivo acutely. This protocol describes an optogenetic system that rapidly inhibits Rho1-mediated actomyosin contractility in Drosophila embryos, revealing the immediate loss of epithelial tension after the inactivation of actomyosin in vivo.

Contractile forces generated by actin and non-muscle myosin II ("actomyosin contractility") are critical for morphological changes of cells and tissues at ...

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

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

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

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

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

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