Name: Collection of Drosophila Embryos and Preparing Them for Optogenetic Stimulation
Uploaded: 2023-04-14T14:50:00
Description: Watch this Scientific Journal Video about Collection of Drosophila Embryos and Preparing Them for Optogenetic Stimulation at JoVE.com

collection of drosophila embryos and preparing them for optogenetic stimulation

Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility in &lt;em&gt;Drosophila&lt;/em&gt;

Actomyosin contractility, the contractile force exerted by non-muscle myosin II (hereafter &#39;myosin&#39;) on the F-actin network, is one of the most important forces in changing cell shape and driving tissue-level morphogenesis1,2. For example, the activation of actomyosin contractility at the apical domain of the epithelial cells results in apical constriction, which facilitates a variety of morphogenetic processes, including epithelial folding, cell extrusion, delamination, and wound healing3,4,5,6,7. The activation of myosin requires phosphorylation of its regulatory light chain. This modification alleviates the inhibitory conformation of the myosin molecules, allowing them to form bipolar myosin filament bundles with multiple head domains on both ends. The bipolar myosin filaments drive the anti-parallel movement of actin filaments and result in the generation of contractile force1,8,9.
The evolutionarily conserved Rho family&#160;small GTPase RhoA (Rho1 in Drosophila)&#160;plays a central role in the activation of actomyosin contractility in various cellular contexts10,11. Rho1 functions as a bimolecular switch by binding either GTP (active form) or GDP (inactive form)12. The cycling between GTP- or GDP-bound Rho1 is regulated by its GTPase-activating proteins (GAPs) and guanine nucleotide-exchange factors (GEFs)13. GEFs function to facilitate the exchange of GDP for GTP and thus activate Rho1 activity. GAPs, on the other hand, enhance the GTPase activity of Rho1 and thus deactivate Rho1. Activated Rho1 promotes actomyosin contractility through interacting with and activating its downstream effectors, Rho-associated kinase (Rok) and Diaphanous14. Rok induces myosin activation and actomyosin contractility by phosphorylating the regulatory light chain of myosin15. In addition, Rok also inhibits the myosin regulatory light chain phosphatase, and hence further promotes myosin filament assembly16. Rok can also phosphorylate LIM kinases, which, when activated, prevent actin disassembly by phosphorylating and inhibiting the actin-depolymerization factor cofilin17,18. Diaphanous is a formin family actin nucleator that promotes actin polymerization, providing a base for myosin to interact with19,20,21.
While the cellular mechanisms that activate actomyosin contractility have been well elucidated, our understanding of its function in regulating dynamic tissue remodeling remains incomplete. Filling this knowledge gap requires approaches that can rapidly inactivate actomyosin at specific tissue regions in vivo and record the immediate impact on tissue behavior and properties. This protocol describes the use of an optogenetic approach to acutely inhibit actomyosin contractility during Drosophila mesoderm invagination, followed by measurement of epithelial tension using laser ablation. During Drosophila gastrulation, the ventrally localized mesoderm precursor cells undergo apical constriction and invaginate from the surface of the embryo by forming an anterior-posteriorly oriented furrow22,23. The formation of ventral furrows has long been used as a model for studying the mechanism of epithelial folding. Ventral furrow formation is administered by the dorsal-ventral patterning system in Drosophila24,25,26,27. The expression of two transcription factors, Twist and Snail, located at the ventral side of the embryo, controls ventral furrow formation and specifies mesodermal cell fate28. Twist and Snail activate the recruitment of the Rho1 GEF RhoGEF2 to the apex of the mesoderm precursor cells via a G-protein coupled receptor pathway and a RhoGEF2 adaptor protein, T4829,30,31,32,33. Next, RhoGEF2 activates myosin throughout the apical surface of the potential mesoderm through the Rho-Rho kinase pathway34,35,36,37,38,39. Activated myosin forms a supracellular actomyosin network throughout the apical surface of the mesoderm primordium, the contractions of which drive apical constriction and result in a rapid increase in apical tissue tension14,37,40.
The optogenetic tool described in this protocol, Opto-Rho1DN, inhibits actomyosin contractility through blue-light dependent plasma membrane recruitment of a dominant negative form of Rho1 (Rho1DN)41. A T19N mutation in Rho1DN eliminates the ability of the mutant protein to exchange GDP for GTP and thus renders the protein perpetually inactive34. A subsequent mutation in Rho1DN, C189Y, eliminates its naive membrane targeting signal42,43. When Rho1DN is infused to the plasma membrane, it binds to and impounds Rho1 GEFs, thereby blocking the activation of Rho1 as well as Rho1-mediated activation of myosin and actin34,44. The plasma membrane recruitment of Rho1DN is achieved through a light-dependent dimerization module derived from Cryptochrome 2 and its binding partner CIB1. Cryptochrome 2 is a blue-light activated Cryptochrome photoreceptor in Arabidopsis thaliana45. Cryptochrome 2 binds to CIB1, a basic helix-loop-helix protein, only in its photoexcited state45. It was later found that the conserved N-terminal, photolyase homology region (PHR), from Cryptochrome 2 (CRY2PHR, hereafter referred to as CRY2) and the N-terminal domain (aa 1-170) of CIB1 (hereafter CIBN) are important for light-induced dimerization46. Opto-Rho1DN contains two components. The first component is the CIBN protein fused with a CAAX anchor, which localizes the protein to the plasma membrane47. The second component is mCherry-tagged CRY2 fused with Rho1DN41. In the absence of blue light, CRY2-Rho1DN remains in the cytoplasm. Upon blue light stimulation, CRY2-Rho1DN is targeted to the plasma membrane through the interaction between membrane-anchored CIBN and excited CRY2. Opto-Rho1DN can be activated by ultraviolet A (UVA) light and blue light (400-500 nm, peak activation at 450-488 nm), or by an 830-980 nm pulsed laser when performing two-photon stimulation41,46,47,48. Therefore, Opto-Rho1DN is stimulated by wavelengths normally used for exciting GFP (488 nm for single photon imaging and 920 nm for two-photon imaging). In contrast, wavelengths commonly used for exciting mCherry (561 nm for single photon imaging and 1,040 nm for two-photon imaging) do not stimulate the optogenetic module and therefore can be used for pre-stimulation imaging. The protocol describes the approaches used to minimize the risk of unwanted stimulation during sample manipulation.
Laser ablation has been extensively employed to detect and measure tension in cells and tissues49. Previous studies have shown that, when laser intensity is appropriately controlled, two-photon laser ablation employing a femtosecond near-infrared laser can physically impair some subcellular structures (e.g., cortical actomyosin networks) without causing plasma membrane rapture50,51. If the tissue is under tension, laser ablation of a region of interest within the tissue results in an immediate outward recoil of the cells adjacent to the ablated region. The recoil velocity is a function of the magnitude of the tension and the viscosity of the media (cytoplasm) surrounding the structures undergoing recoil49. Because of the superior penetration depth of the near-infrared lasers and the ability to achieve well-confined focal ablation, two-photon laser ablation is particularly useful for detecting tissue tension in vivo. As demonstrated in this protocol, this method can be easily combined with Opto-Rho1DN-mediated inactivation of actomyosin contractility to investigate the direct impact of Rho1-dependent cellular contractility on tissue mechanics during dynamic tissue remodeling.

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

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

Our research studies tissue morphogenesis, the formation of complex three-dimensional tissue structures in development. We are interested in the genes and the molecules that regulate morphogenesis and seek to understand the physical principles underlying morphogenesis. For example, how mechanical forces are generated and how they drive tissue rehabilitation.Contractile forces generated by filamentous actin and non-muscle myosin II, also known as actomyosin contractility, is one of the most important forces that drive tissue morphogenesis. Our current research addresses how actomyosin contractility mediates the folding of blood epithelial cell sheets, a fundamental tissue construction mechanism in development. An in-depth understanding of the role of actomyosin contractility in epithelial folding and other morphogenetic processes requires approaches that can quickly inactivate actomyosin at designated time and location, and records the immediate impact of tissue behavior and properties.However, this is difficult to achieve using conventional genetic approaches. The approach described in this protocol is designed to address how cells and tissues respond to sudden changes in mechanical forces that normally drive tissue remodeling. This is an important question but has been difficult to tackle due to limited approaches that can quickly alter mechanical forces in intact tissues.In this study, we developed an optogenetic tool to inactivate actomyosin at specific tissue regions in Drosophila embryos quickly. We found that combining these two with laser ablation to investigate the direct impact of actomyosin contractility on tissue mechanics in epithelial folding. Our findings provide new insights into the mechanical mechanism apical constriction media to epithelial folding.With manual modifications, the protocol can be easily adapted to study the function of actomyosin contractility in a wide range of morphogenetic processes.

Watch this Scientific Journal Video about Collection of Drosophila Embryos and Preparing Them for Optogenetic Stimulation at JoVE.com

Collection of Drosophila Embryos and Preparing Them for Optogenetic Stimulation  

Collection of Drosophila Embryos and Preparing Them for Optogenetic Stimulation

To begin collecting the drosophila embryos, change the apple juice plate from the cup and label the plate. Cover the surface of the plate with a thin layer of halocarbon oil 27. Next position, an orange red plastic shield on the stage of an upright stereoscope.Place the apple juice plate on the orange red shield. Turn on the transmitted light of the stereo microscope to illuminate the sample. Collect 5 to 15 embryos from the apple juice plate using a pair of tweezers.Gently blot the embryos on a paper towel, sized approximately 1.5 by 1.5 centimeters. Then add several drops of freshly prepared 40%bleach to a new paper towel using plastic transfer pipette to cover the paper towel with a thin layer of bleach. Transfer the embryo from the dry paper towel to the bleach soaked paper towel and ensure the embryos are soaked in the bleach.Wait two minutes for the embryo to become dechorionated. After dechorionation, using tweezers, blot the paper towel on a large piece of tissue paper to remove the excess bleach. Rinse the embryos with deionized water eight times to remove residue bleach.Using an eyelash tool, transfer the embryo from the paper towel to a 35 millimeter glass bottom dish. Then add deionized water to the dish to cover the embryos completely. Finally, fine tune the position and orientation of the embryos using the eyelash tool.Place the dish with the embryos inside a light proof black box to protect the sample from light exposure during the transfer process.

collection-drosophila-embryos-preparing-them-for-optogenetic

Research

JoVE Journal

Developmental Biology

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.

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 ...

Optogenetic Stimulation, Laser Ablation, and Imaging of Drosophila Embryos

Optogenetic Stimulation, Laser Ablation, and Imaging of Drosophila Embryos  

To begin, turn off the room light. Select Ocular under the ocular panel in the FlowView software. Change the cube turret to four TRITC, and turn off the touch panel controller by clicking off under backlight of touch panel controller in the software.Then turn off the computer screen. Open the front side of the black cloth cover on the microscope. Take the 35 millimeter glass-bottomed dish containing the embryos from the black box and place it on the microscope stage.Then turn on the fluorescence illumination unit, and use the eyepiece to identify the embryo of interest. Bring it into focus. Close the black cloth cover to protect the sample from light.Turn on the computer screen to access the software that controls the microscope. Change the ocular to LSM in the software for image acquisition. Perform laser ablation in control unstimulated embryos using a 25 times magnification water immersion objective.Click Bright Z, Sequence Manager, and LSM stimulation from the tool window. Set the scanner type as Galvano, and the scan size as 512 by 512. Turn on channel one and channel three under the PMT setting panel to allow the use of the 1040 nanometer laser, and click live four times speed to visualize the embryo.Rotate the embryo using the rotation function to make the anterior posterior axis vertically oriented, and set the zoom to three. Draw a region of interest or ROI using the shape tool under scan settings, and set the ROI size in the reference panel. Next, set the ROI as 512 pixels in width and 100 pixels in height.To set the acquisition parameters for the pre-ablation Z stack, register the embryo's surface as zero under the Z section. Set the start as zero and the end as 100 micrometers. Set the step size as two micrometers, and activate the Z acquisition mode by checking Z under the series tab.Using the bright Z function, set the 1040 nanometer laser intensity to increase linearly from 3%to 7%Save the current imaging setting as the first task of the pipeline by clicking LSM in Sequence Manager. To set acquisition parameters for the pre-ablation movie, set an ROI of 512 by 512 pixels near the embryo's ventral surface as previously demonstrated. Set the 1, 040 nanometer laser intensity to 3%Check time, and uncheck Z under the series panel.Keep the interval as free run under the time lapse panel, and set the cycle as 10. Save the current setting as the next task of the pipeline by clicking LSM in Sequence Manager. To set the parameters for laser ablation, define a 3D region immediately below the vitelline membrane.Set the start of the Z stack as the plane, and the end as 20 micrometers deeper. Set the step size as 1.5 micrometers. Turn on channel two and channel four under the PMT setting panel to allow the use of the 920 nanometer laser.Set the intensity of the laser to 30%and set image acquisition with the laser for a single Z stack within the defined 3D region. Save the current setting as the next task of the pipeline by clicking LSM in Sequence Manager. To set acquisition parameters for the post ablation movie, set image acquisition for a 100 frame single Z plane post ablation movie using the 1, 040 and 920 nanometer lasers.Set the intensity of the lasers to 3%and 0.3%Save the current setting as the next task of the pipeline by clicking LSM in Sequence Manager. Select sequence under acquire. Change the data saving path and file name as needed.Click ready, and wait for the software to initialize the pipeline, then click start to execute the pipeline. to perform laser ablation in stimulated embryos, Set acquisition parameters for the pre-ablation Z stack as demonstrated for the unstimulated embryos. Save the current setting as the first task of the pipeline by clicking LSM in Sequence Manager.To set parameters for optogenetic stimulation within a defined ROI, change the zoom to one, and select an ROI that covers the embryo's ventral surface. Turn off the channel one to channel four detectors, click LSM stimulation, uncheck continuous within duration, and type 12 seconds. Save the current setting as the next task of the pipeline by clicking stimulation in Sequence Manager.Set a three minute wait time after stimulation to ensure total inactivation of myosin and apical F-actin disassembly and achieve a static tissue morphology before laser ablation. Next, set acquisition parameters for the single Z plane pre-ablation movie as demonstrated for the unstimulated embryos, except that the 1, 040 and 920 nanometer lasers are used for image acquisition. Turn on the channel one to channel four detectors.Set the intensity of the lasers to 3%and 0.3%Save the current setting as the next task of the pipeline by clicking LSM in Sequence Manager. Set parameters for laser ablation as demonstrated for the unstimulated embryos. Save the current setting as the next task of the pipeline by clicking LSM in Sequence Manager.Set acquisition parameters for the single Z plane post-ablation movie as demonstrated for the unstimulated embryos. Save the current setting as the next task of the pipeline by clicking LSM in Sequence Manager. Select sequence under acquire.Change the data saving path and file name as needed. Click ready and wait for the software to initialize the pipeline, then click start to execute the pipeline. In the unstimulated embryos undergoing apical constriction, spaghetti squash mCherry became enriched at the medial apical region, whereas CRY2Rho one dominant negative mCherry was cytosolic.In the stimulated embryos, the CRY2Rho one dominant negative mCherry signal became plasma membrane localized, whereas the medial apical signal of spaghetti squash mCherry completely disappeared. Laser ablation of the unstimulated embryos within the constriction domain led to a rapid tissue recoil along the anterior posterior axis, whereas laser ablation in the stimulated embryos did not result in noticeable tissue recoil. The ablation was quantified and shown here.