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Live Imaging of Myogenesis in Indirect Flight Muscles in Drosophila
果蝇间接飞行肌中肌生成的实时成像   

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Abstract

The indirect flight muscles (IFMs) are the largest muscles in the fly, making up the bulk of the adult thorax. IFMs in Drosophila are generated during pupariation by fusion of hundreds of muscle precursor cells (myoblasts) with larval muscle templates (myotubes). Prominent features, including the large number of fusion events, the structural similarity to vertebrate muscles, and the amenability to the powerful genetic techniques of the Drosophila system make the IFMs an attractive system to study muscle cell fusion. Here we describe methods for live imaging of IFMs, both in intact pupae, and in isolated IFMs ex-vivo. The protocols elaborated upon here were used in the manuscript by (Segal et al., 2016).

Keywords: Myoblast fusion(成肌细胞融合), Live imaging(实时成像), Indirect flight muscle(间接飞行肌), Drosophila(果蝇), Muscle(肌肉), ex-vivo culture(离体培养)

Background

While Drosophila embryonic muscles have long been an established model system for the study of muscle development (Volk, 1999; Chen and Olson, 2004; Abmayr et al., 2008; Richardson et al., 2008) the adult Drosophila indirect flight muscles (IFMs), which form during pupal stages, have emerged in recent years as a complementary system to address cell-biological processes during myogenesis (Dutta, 2006; Oas et al., 2014; Weitkunat et al., 2014; Shwartz et al., 2016). Their large size, ample fusion events, structural similarity to vertebrate muscles, and amenability to powerful genetic techniques of the Drosophila system make the IFMs an attractive system to study muscle development. Historically, study of IFM development has been limited compared to embryonic muscle for several reasons. First, classic genetic approaches are difficult to implement in IFMs due to functional requirements earlier in development for many of the genes potentially involved in muscle development in the adult and to the syncytial nature of muscles, which restricts the usefulness of clonal analysis. Relatively recent advances in the available tools utilizing the GAL4-UAS system allow circumvention of these limitations, by expressing RNAi in a tissue specific manner. The power of this approach was demonstrated in a comprehensive screen for genes involved in pupal myogenesis (Schnorrer et al., 2010), and has been successfully implemented in several recent studies of myoblast fusion in IFMs (Mukherjee et al., 2011; Gildor et al., 2012; Dhanyasi et al., 2015; Segal et al., 2016). In addition, the technical challenges associated with dissection, accessibility, and visualization of IFMs have been overcome by advances in techniques and technology (Weitkunat and Schnorrer, 2014; Segal et al., 2016). This protocol is an expanded version of the methods used in the manuscript by (Segal et al., 2016), and is intended to contribute to the growing repertoire of techniques for study of IFMs. Here we describe methods for live imaging of IFMs, both in intact pupae, and in isolated IFMs ex-vivo. While previous work focused on stages of myotube growth via fusion (18-22 h after puparium formation [APF], 25 °C), these methods are readily applicable to other stages of IFM myogenesis, starting at 12 h APF onwards. Imaging of intact pupae can be suitable for studies of developmental processes which span several hours, while imaging of ex-vivo cultures is intended to better visualize finer structural details and dynamic behaviors over shorter time periods (e.g., 1 h). Parts of this protocol are variations on (Weitkunat and Schnorrer, 2014).

Materials and Reagents

  1. Glass single frosted microscope slides (Thermo Fisher Scientific, catalog number: 421-004T )
  2. Cover slip
  3. Custom-made plexiglass slide with round opening in center (will serve as basis for viewing chamber)
    Note: The opening should be 1.25 cm in diameter.
  4. Double-sided tape
  5. Kim-wipes
  6. Toothpick
  7. 8-well chamber slide (μ-slide) (ibidi, catalog number: 80826 )
  8. 200 μl and 1,000 μl pipette tips
  9. Sylgard silicone plates, stained with charcoal (Dow Corning, catalog number: 3097358-1004 )
  10. 1 cm Minutian pins for silicon plate (Fine Science Tools, catalog number: 26002-10 )
  11. Flies expressing muscle-specific fluorescent markers (e.g., mef2-GAL4> UAS-CD8-GFP)
  12. Schneider’s medium
  13. Fetal bovine serum (FBS)
  14. BD Matrigel matrix growth factor, reduced (Corning, catalog number: 354230 )
  15. Halocarbon oil

Equipment

  1. Thin paint brush
  2. Forceps (Dumont 55) (Fine Science Tools, catalog number: 11255-20 )
  3. Vannas Spring scissors, 2.5 mm Cutting Edge (Fine Science Tools, catalog number: 15001-08 )
  4. Confocal microscopy system (e.g., ZEISS, model: LSM 780 ) equipped with an inverted microscope and a 40x water immersion objective
  5. 200 μl and 1,000 μl pipette
  6. 125 ml Erlenmeyer flask
  7. 25 °C incubator to grow flies (Elcon company, custom-made)
  8. Vials with fly food to raise and collect flies
  9. 37 °C incubator for matrigel solidification (Thermo Fisher Scientific, Thermo ScientificTM, model: HeracellTM 150i )
  10. Oxygen tank with attached rubber tube
  11. Binocular dissecting microscope (Nikon Instruments, model: SMZ645 )
  12. Fluorescent binocular microscope (Leica, model: Leica MZ16F )

Software

  1. Zen Black software (ZEISS)
  2. Microsoft Excel
  3. R Statistical software (can be downloaded at: https://www.r-project.org/)

Procedure

  1. Preparation of intact pupae for live imaging
    Note: IFM myoblast fusion is at its peak at 18-22 h APF when grown at 25 °C, but this method can be used to image IFMs before or after that time range. Pupae at 12 h APF or earlier are more difficult to work with, since head eversion has yet to occur, so the pupal case is still attached to the pupa.
    1. Select fly line with IFMs expressing a fluorescent marker. Pupae are grown to the desired stage, by collecting white pre-pupae (which are 0-1 h old) into a new vial, and growing at 25 °C (e.g., for 18-20 h). Pre-pupae can be identified by their formed antennae and lack of any pigmentation (white color).
    2. Prepare viewing chamber, by attaching a cover slip to the custom-made plexiglass slide with opening, with tape on either side of the opening (so that the coverslip is positioned over the opening) (Figure 1A).


      Figure 1. Intact pupae. A. Custom-made slide for imaging of intact pupae. Coverslip is taped from both sides over opening. B. Pupa adhered to double-sided tape on glass slide. For dissection, first remove the ‘lid’ of the pupal case with forceps (1). Then, place one side of the forceps between the pupa and the pupal case, and move downward and outward to tear the pupal case open (2). Lift the formed ‘flap’ to other side to expose pupa (3). C. Pupal case is peeled from side and opened to double-sided tape beneath to expose pupa. D. GFP fluorescent filter view, via a dissection microscope and through a cover-slip, showing the dorsal side of an intact pupa, immersed in halocarbon oil. Boxed area enlarged on bottom right showing the 6 IFM fibers. Scale bars = 500 μm (B and C), 50 μm (D).

    3. On a glass microscope slide, place a strip of double-sided tape.
    4. Carefully, lift staged pupae with brush from vial and place them onto a Kim-wipe. Carefully brush pupae against Kim-wipe to remove possible food residue.
    5. Using brush, place a pupa dorsal side up (flat side down) on the double-sided tape strip. Press down lightly on the pupa with a brush to ensure it is well adhered, without injuring the pupa.
    6. The next few steps are designed to break open the pupal case and remove/transfer the intact pupa to the viewing chamber. At these developmental stages head eversion should be complete, so that a gap exists between the pupal case and the head. With forceps, carefully capture the extreme anterior end of the pupal case, and lift upward to peel off the circular ‘lid’ of the pupal case (Figure 1B).
    7. Slide forceps gently into the anterior opening you created, so that one arm of forceps protrudes into the space between the pupa and the pupal case.
    8. Pull outward to tear the pupal case. Continue this movement until the pupal case is opened along its dorsal aspect all the way to the posterior end.
    9. The ‘flap’ that has formed on the upper portion of the pupal case can now be lifted/opened by the forceps and attached to the tape on the opposite side, thereby revealing the pupa (Figures 1B and 1C).
    10. Place the closed forceps under the pupa, and gently lift and transfer it to the cover slip attached to custom-made slide. The pupa should stick to the forceps. If not, lift by the forming legs of the pupa, underneath the abdomen.
    11. Using the forceps, orient the pupa so that the thorax (dorsal side) faces the cover-slip.
    12. Place a drop of halocarbon oil on a separate glass slide. From this drop, using a toothpick, pick up a smaller drop of oil and gently transfer onto the pupa, so that it is mostly immersed in oil. Put only enough oil to cover the pupa, as excessive oil can cause movement of the pupa during long-term imaging. Absorb excess oil with a Kim-wipe (Figure 1C).
    13. Orient the pupa so that it is slightly tilted to one side at about a 45° angle, for a better angle of imaging for one of the IFMs.
    14. Image slide immediately using the inverted confocal microscope system. Live intact pupae have been successfully imaged for up to 12 h. See more details for image acquisition below.

  2. Dissection and isolation of IFMs for ex-vivo live imaging
    1. Prepare an ibidi μ-slide coated with Matrigel as follows. Matrigel should be diluted 1:10 in PBS from stock and thawed at 4 °C overnight, as described by manufacturer here. With a micropipette, transfer 100 μl into a single well, and spread with tip so that it covers the entire surface of the well. Re-pipette and remove the entire 100 μl Matrigel. The small amount that necessarily remains will create the Matrigel coat. For preparation of multiple wells, the same 100 μl can be re-used. Discard remaining liquid. Place ibidi μ-slide at 37 °C for 30 min in order to allow the Matrigel to harden.
    2. Prepare medium for live imaging. First, oxygenate Schneider’s medium as follows. Place 20-50 ml Schneider’s medium in 125 ml Erlenmeyer flask. Attach 1,000 μl micropipette tip to the end of a tube attached to an oxygen tank. Tilt the flask slightly (e.g., by placing bottom on another micropipette tip, see Figure 2). Place tube so that the tip is immersed in the medium, and gradually increase oxygen pressure until bubbles are seen. Oxygenate for 10 min. After oxygenating, add 10% FBS to desired amount of medium. Adding FBS before oxygenating is not recommended as it could create foam.


      Figure 2. Oxygenation of Schneider’s medium. See text for details. In this set-up, the tube is taped to the wall for convenience of placement.

    3. Stage and isolate a pupa from its pupal case, as in steps A1-A10 above. Transfer the pupa to a darkened Sylgard silicon plate, dorsal (curved) side down. Place 200 μl Schneider’s medium + 10% FBS, oxygenated, on pupa.
    4. Insert pin between head and thoracic segments of the pupa, angling ~135° away from thoracic segment, so as to not hinder the view of the thorax. Pin should be sharp to smoothly puncture the pupa (Figure 3A).


      Figure 3. Isolation of IFMs for ex-vivo live imaging. A. Pinned pupa. First, cut posterior tip of abdomen (1). Then, cut ventro-lateral cuticle on either side (2 and 3), and finally cut horizontally to remove ventral cuticle (4). (B-B’) Opened pupa before removing fat and other irrelevant tissue viewed by transmitted (B) and flourescent (B’) light. Note that IFMs cannot be seen at this stage, as they are lightly adhered to the dorsal cuticle and are obscured by surrounding tissue. (C-C’) Pupa after clearing of other tissues viewed by transmitted and fluorescent light (C) or fluorescent light alone (C’). IFMs are circled. (D-D’) Isolated IFM pairs viewed by transmitted and fluorescent light (D) or fluorescent light alone (D’). Scale bars = 200 μm (A), 50 μm (D).

    5. Under binocular, with microdissection scissors, snip off the posterior tip of the abdominal cuticle.
    6. With scissors, carefully slide up and cut the lateral sides of the cuticle, on both sides. Cut across the pupa below pin, allowing to remove and discard the ventral portion of the cuticle.
    7. Under fluorescent binocular, place blue light filter to visualize GFP. With forceps, gently remove fat bodies, intestines, and other tissues, avoiding contact with the dorsal cuticle and attached tissues. At these stages, the IFMs should be lightly adhered to the dorsal cuticle (Figures 3B and 3B’). Avoid clearing trachea at this stage.
    8. Depending on the specific fluorescent line used, there may be other tissues that also express GFP. At these stages, the IFMs should be recognizable morphologically in addition to GFP fluorescence, as 6 elongated fibers, lightly attached to both sides of the dorso-lateral cuticle, in the mid-anterior portion of the thoracic segment. Visualizing the IFMs without GFP fluorescence is more challenging.
    9. Gently clear remaining fat and trachea with forceps to find and expose the IFMs. In some cases, IFMs are lightly attached to trachea and must be separated with forceps (Figures 3C and 3C’, see ‘Notes’ section for some additional tips).
    10. Once IFMs are exposed, pinch with forceps underneath IFMs to detach them from the cuticle.
    11. Isolate the IFMs by gently pushing them with side of forceps to a region of the medium that is outside the dissected pupa (Figures 3D and 3D’).
    12. Transfer 200 μl Schneider’s medium + 10% FBS, oxygenated to a Matrigel-coated well in ibidi plate.
    13. With micropipette, wet 200 μl micropipette tip with clean medium. In the silicon plate, lift a small amount of liquid into the tip, and with micropipette still pressed, lift up the isolated IFMs into 50 μl liquid.
    14. Gently transfer the IFMs and liquid into the medium-filled well. The IFMs should sink to the surface of the well.
    15. With side of forceps, gently press, flatten, and orient the IFMs for imaging in the ibidi slide well.
      Note: Optionally, chemical inhibitors (e.g., LatrunculinA) or dyes (e.g., SiR Tubulin) may be added to the medium at relevant concentrations.
    16. Take immediately for imaging using the confocal microscope system (e.g., ZEISS 780). Isolated IFMs can be imaged ex-vivo for up to 1.5 h. Time-lapse movies were taken at 40x water immersion objective, optimal for thick biological samples. The entire muscle is approximately 35 μm thick. Analysis was often done on maximum intensity projections of acquired z-stacks or subsets of z-stacks, using the Zen software (ZEISS). The following acquisition settings allowed imaging most of the depth of the IFMs at a time resolution of approximately 1.5 min per stack: 33 slices of 1.1 μm thickness, 512 x 512 resolution, line averaging of 4, 2x digital zoom. For better time resolution, decrease averaging and/or number of slices.

Data analysis

In general, visualization over time of fluorescent markers of interest can provide insight of dynamic behavior simply by observation of time-lapse of the acquired data. Time-lapse movies of single slices or maximum intensity projections of the data were created using Zen software (ZEISS). For example, a time-lapse movie of a single optical slice from an ex-vivo culture of dissected IFMs expressing Lifeact-GFP to label actin fibers (Video 1) displays myoblasts with actin-based protrusions emanating from their edges, which appear to be entangled within one another. The myoblasts are surprisingly immobile over time.


ly:Arial;font-size:10pt;margin-left:20px;"> Video 1. Ex-vivo IFM expressing mef2 > Lifeact-GFP. Movie shows a single slice of outer surface of isolated IFM, displaying the many myoblasts associated to the myotube.

Quantification of length, frequency, etc. of specific features may also be performed in this system. For example, Segal et al. 2016 measured the lengths of filopodia in IFMs expressing the membrane marker Gap-GFP (Ritzenthaler et al., 2000). In this case, dynamics over time were not necessary, so a z-stack at a single timepoint was acquired (Figure 4). Using Zen Black software, a subset of z-slices were selected to include the filopodia of the myotube but exclude background from neighboring myotubes. Then, a maximum intensity projection (MIP) of the subset of slices was made. An area of interest was selected and cropped (Figure 4, red box), and brightness was adjusted to highlight the filopodia. The cropped raw data was transferred to Zen Blue software, for measurement of filopodia lengths. The output of lengths of filopodia was transferred to Excel for analysis. Statistical analysis of differences in filopodia length between experimental groups was done in R software, but could be done on any statistical software.


Figure 4. Analysis of filopodia length as in Segal et al., 2016. See text for details.

Notes

  1. Dorso-longitudinal muscles (DLMs) and dorso-ventral muscles (DVMs), the two types of IFMs, remain attached. DLMs can be distinguished as the larger of the two muscle groups. Previous work imaged the DLMs specifically (Segal et al., 2016).
  2. During development the DLMs split from three fibers to 6 fibers, between 16-18 h APF. Therefore, IFMs may be in the midst of splitting if dissected during this time range.
  3. Ideally, an intact DLM will include 3 or 6 fibers (see above) and the surrounding myoblasts. However, it is not uncommon for individual myofibers to be lost during dissection. In general, myoblasts will still remain associated with the remaining myofibers, and these samples can still be imaged, as long as the individual myofibers remain intact.

Acknowledgments

This work was done under the supervision of and with guidance from Prof. Benny Shilo and Dr. Eyal Schejter, at Weizmann Institute of Science in Rehovot, Israel. This work was supported by a grant from the Israel Science Foundation. Parts of this protocol were adapted from (Weitkunat and Schnorrer, 2014). This protocol was used in (Segal et al., 2016).

References

  1. Abmayr, S. M., Zhuang, S. and Geisbrecht, E. R. (2008). Myoblast fusion in Drosophila. Methods Mol Biol 475: 75-97.
  2. Chen, E. H. and Olson, E. N. (2004). Towards a molecular pathway for myoblast fusion in Drosophila. Trends Cell Biol 14(8): 452-460.
  3. Dhanyasi, N., Segal, D., Shimoni, E., Shinder, V., Shilo, B. Z., VijayRaghavan, K. and Schejter, E. D. (2015). Surface apposition and multiple cell contacts promote myoblast fusion in Drosophila flight muscles. J Cell Biol 211(1): 191-203.
  4. Dutta, D. and VaijayRaghavan, K. (2006). Metamorphosis and the formation of the adult musculature. In: Muscle Development in Drosophila. Springer 125-142.
  5. Gildor, B., Schejter, E. D. and Shilo, B. Z. (2012). Bidirectional Notch activation represses fusion competence in swarming adult Drosophila myoblasts. Development 139(21): 4040-4050.
  6. Mukherjee, P., Gildor, B., Shilo, B. Z., VijayRaghavan, K. and Schejter, E. D. (2011). The actin nucleator WASp is required for myoblast fusion during adult Drosophila myogenesis. Development 138(11): 2347-2357.
  7. Oas, S. T., Bryantsev, A. L. and Cripps, R. M. (2014). Arrest is a regulator of fiber-specific alternative splicing in the indirect flight muscles of Drosophila. J Cell Biol 206(7): 895-908.
  8. Richardson, B., Beckett, K. and Baylies, M. (2008). Visualizing new dimensions in Drosophila myoblast fusion. Bioessays 30(5): 423-431.
  9. Ritzenthaler, S., Suzuki, E. and Chiba, A. (2000). Postsynaptic filopodia in muscle cells interact with innervating motoneuron axons. Nat Neurosci 3(10): 1012-1017.
  10. Schnorrer, F., Schonbauer, C., Langer, C. C., Dietzl, G., Novatchkova, M., Schernhuber, K., Fellner, M., Azaryan, A., Radolf, M., Stark, A., Keleman, K. and Dickson, B. J. (2010). Systematic genetic analysis of muscle morphogenesis and function in Drosophila. Nature 464(7286): 287-291.
  11. Segal, D., Dhanyasi, N., Schejter, E. D. and Shilo, B. Z. (2016). Adhesion and fusion of muscle cells are promoted by filopodia. Dev Cell 38(3): 291-304.
  12. Shwartz, A., Dhanyasi, N., Schejter, E. D. and Shilo, B. Z. (2016). The Drosophila formin Fhos is a primary mediator of sarcomeric thin-filament array assembly. Elife 5.
  13. Volk, T. (1999). Singling out Drosophila tendon cells: a dialogue between two distinct cell types. Trends Genet 15(11): 448-453.
  14. Weitkunat, M., Kaya-Copur, A., Grill, S. W. and Schnorrer, F. (2014). Tension and force-resistant attachment are essential for myofibrillogenesis in Drosophila flight muscle. Curr Biol 24(7): 705-716.
  15. Weitkunat, M. and Schnorrer, F. (2014). A guide to study Drosophila muscle biology. Methods 68(1): 2-14.

简介

间接飞行肌肉(IFM)是飞行中最大的肌肉,构成成年胸部的大部分。 通过将数百种肌肉前体细胞(成肌细胞)与幼虫肌肉模板(肌管)融合,在果蝇中产生IFMs 。 突出的特征,包括大量的融合事件,与脊椎动物肌肉的结构相似性,以及对果蝇系统强大的遗传技术的适应性使得IFM成为研究肌肉细胞融合的有吸引力的系统。 在这里,我们描述了在完整的蛹中和在离体的独立的IFM中实时成像IFM的方法。 (Segal等人,2016年)在手稿中使用了这里阐述的方案。
【背景】胚胎肌肉长期以来一直是肌肉发育研究的建立模型系统(Volk,1999; Chen和Olson,2004; Abmayr等人,2008; Richardson pup biological(Dutta,2006; Oas等人,2014; Weitkunat等人,2014; Shwartz等人,2016) 。它们的大尺寸,丰富的融合事件,与脊椎动物肌肉的结构相似性,以及对果蝇系统强大的遗传技术的适应性使得IFM成为研究肌肉发育的有吸引力的系统。历史上,与胚胎肌相比,IFM发展的研究受到限制,原因有几个。首先,由于许多可能参与成年肌肉发育的基因和肌肉合成性质的基因的早期功能需求,IFM中难以实施经典遗传学方法,这限制了克隆分析的有用性。使用GAL4-UAS系统的可用工具的相对最近进展通过以组织特异性方式表达RNAi来避免这些限制。这种方法的力量在综合筛查蛹蜕膜基因(Schnorrer等人,2010)中得到证实,并且已经在IFMs中成肌细胞融合的最近几项研究中成功实施(Mukherjee et al。,2011; Gildor等人,2012; Dhanyasi等人,2015; Segal等人。,2016)。此外,IFM的解剖,可及性和可视化相关的技术挑战已经通过技术和技术的进步而得到克服(Weitkunat和Schnorrer,2014; Segal等人,2016)。该协议是由(Segal等人,2016)手稿中使用的方法的扩展版本,旨在促进研究IFM的技术越来越多。在这里,我们描述了在完整的蛹中和在离体的独立的IFM中实时成像IFM的方法。虽然以前的工作集中在通过融合进行肌管生长的阶段(瞳孔形成后18-22小时[APF],25°C)),但是这些方法很容易适用于从12小时APF开始的IFM肌生成的其他阶段。完整蛹的成像可适用于跨越数小时的发育过程的研究,而离体培养物的成像旨在更好地在更短的时间段内可视化更精细的结构细节和动态行为(例如,1小时)。该协议的一部分是(Weitkunat和Schnorrer,2014)的变体。

关键字:成肌细胞融合, 实时成像, 间接飞行肌, 果蝇, 肌肉, 离体培养

材料和试剂

  1. 玻璃单个磨砂显微镜载玻片(Thermo Fisher Scientific,目录号:421-004T)
  2. 封面
  3. 定制的有机玻璃滑块与圆形开口在中心(将作为观察室的基础)
    注意:开口直径应为1.25厘米。
  4. 双面胶带
  5. 金擦拭物
  6. 牙签
  7. 8孔室玻片(μ-slide)(ibidi,目录号:80826)
  8. 200μl和1,000μl移液管吸头
  9. Sylgard硅胶板,用木炭染色(道康宁,目录号:3097358-1004)
  10. 1厘米硅片的微型针脚(精细科学工具,目录号:26002-10)
  11. 表达肌肉特异性荧光标记的苍蝇(例如,,mef2 -GAL4> UAS-CD8-GFP)
  12. 施耐德的媒介
  13. 胎牛血清(FBS)
  14. BD Matrigel基质生长因子减少(Corning,目录号:354230)
  15. 卤碳油

设备

  1. 薄油漆刷
  2. 镊子(Dumont 55)(精细科学工具,目录号:11255-20)
  3. Vannas弹簧剪刀,2.5毫米刀刃(精细科学工具,目录号:15001-08)
  4. 配有倒置显微镜和40x水浸物镜的共聚焦显微镜系统(例如,ZEISS,型号:LSM 780)
  5. 200μl和1,000μl移液器
  6. 125ml锥形瓶
  7. 25°C孵化器种植苍蝇(Elcon公司,定制)
  8. 有飞行食物的小瓶可以养殖和收集苍蝇
  9. 37℃用于基质胶凝固的培养箱(Thermo Fisher Scientific,Thermo Scientific TM,型号:Heracell TM 150i)
  10. 带有橡胶管的氧气罐
  11. 双目解剖显微镜(Nikon Instruments,型号:SMZ645)
  12. 荧光双目显微镜(Leica,型号:Leica MZ16F)

软件

  1. 禅黑软件(ZEISS)
  2. Microsoft Excel
  3. R统计软件(可从以下网址下载: https://www.r-project。 org /

程序

  1. 用于实时成像的完整蛹的准备
    注意:当在25°C生长时,IFM成肌细胞融合在18-22 h APF处于峰值,但该方法可用于在该时间范围之前或之后对IFM进行成像。 12小时APF或更早的蛹是更难以合作的,因为头端外翻尚未发生,所以瞳孔病例仍然附着在蛹上。
    1. 选择表达荧光标记的IFM的飞行线。通过将白色蛹前(0-1小时)收集到新的小瓶中,并在25℃(例如)生长18-20小时,将蛹生长至期望的阶段。 。可以通过它们形成的天线和缺乏任何色素沉着(白色)来鉴定蛹前。
    2. 准备观察室,通过将打开的盖板粘贴到定制的有机玻璃滑动件上,使开口两侧的胶带(使盖玻片位于开口上方)(图1A)。


      图1.完整蛹。 A.用于成像完整蛹的定制幻灯片。盖子从开口两侧胶带。 B.傀儡在玻璃片上贴上双面胶带。对于解剖,首先用镊子(1)去除蛹壳的“盖子”。然后,将镊子的一侧放在蛹和蛹壳之间,向下和向外移动以撕开蛹壳(2)。将形成的“襟翼”提升到另一侧以暴露蛹(3)。 C.蛹壳从侧面剥下,打开到双面胶带,露出蛹。 D. GFP荧光过滤器视图,通过解剖显微镜和覆盖滑动,显示完整蛹的背侧,浸入卤碳油中。右下方放大盒装区域,显示6条IFM纤维。刻度棒=500μm(B和C),50μm(D)。

    3. 在玻璃显微镜载玻片上,放置一条双面胶带。
    4. 小心地,用小瓶的刷子提起分阶段的蛹,并将它们放在Kim-wipe上。仔细地揉搓金w擦去可能的食物残渣。
    5. 使用刷子,将蛹背面朝上(平面朝下)放置在双面胶带上。用刷子轻轻按下蛹,以确保其粘附良好,而不会伤害蛹。
    6. 接下来的几个步骤旨在打破蛹壳,并将完整的蛹移除/转移到观察室。在这些发展阶段,头部外翻应该是完整的,所以瞳孔与头部之间存在差距。用镊子仔细捕捉蛹壳的前端,向上抬起蛹壳的圆形“盖子”(图1B)。
    7. 将镊子轻轻地插入您创建的前开口,使镊子的一个臂伸入蛹和蛹壳之间的空间。
    8. 向外拉,撕开蛹壳。继续这个运动,直到瞳孔沿其背部方向一直打开到后端。
    9. 现在可以通过镊子将瞳孔上部形成的“襟翼”抬起/打开,并将其附着在相对侧的胶带上,从而露出蛹(图1B和1C)。
    10. 将封闭的镊子放在蛹下,轻轻提起并将其转移到附在定制滑块上的盖子上。蛹应坚持镊子。如果没有,请通过瞳孔的形成腿抬起腹部。
    11. 使用镊子,定向蛹,使胸部(背侧)面向覆盖滑块。
    12. 将卤代烃油放在单独的玻璃片上。从这一滴,使用牙签,拿起一小滴油,轻轻地转移到蛹上,使其大部分浸在油中。只能用足够的油来覆盖蛹,因为过量的油会导致长期成像期间蛹的运动。用金擦拭物吸收多余的油(图1C)。
    13. 定向蛹,使其以约45°的角度稍微倾斜到一侧,以便为其中一个IFM获得更好的成像角度。
    14. 使用倒置的共聚焦显微镜系统立即图像幻灯片。活体完整的蛹已成功成像长达12小时。请参阅下面的图像采集的更多细节。

  2. 离体和离散IFMs用于离体活体成像
    1. 准备用Matrigel涂布的ibidiμ-滑片,如下。基质胶应在PBS中以1:10稀释,并在4℃下解冻过夜,如制造商所述 here 。使用微量移液管将100μl转移到单个孔中,并用尖端传播,以覆盖整个井的表面。重新吸取和去除整个100μlMatrigel。必须保留的少量将创造Matrigel大衣。为了制备多个孔,可以重复使用相同的100μl。丢弃剩余的液体。将ibidiμ-slide在37℃下放置30分钟,以使Matrigel硬化。
    2. 准备用于实时成像的介质。首先,含氧化合物施耐德的介质如下。将20-50ml施耐德氏培养基置于125ml锥形瓶中。将1,000μl微量移液器末端连接到连接到氧气罐的管道的末端。通过将底部放置在另一个微量移液管尖端上,稍稍倾斜(例如,),见图2)。放置管子使得尖端浸入介质中,并逐渐增加氧气压力,直到看到气泡。氧合10分钟加氧后,加入10%FBS至所需量的培养基。不建议在充氧前添加FBS,因为它可能会产生泡沫。


      图2.施耐德介质的氧化。详见文本。在这种设置中,管子被贴在墙上,以方便放置。

    3. 如同上述步骤A1-A10,将蛹与其蛹情况相分离并分离。将蛹转移到黑暗的Sylgard硅板上,背面(弯曲)侧向下。在蛹上放置200μlSchneider's培养基+ 10%FBS,充氧。
    4. 在蛹的头部和胸部段之间插入针头,与胸廓段倾斜约135°,以免阻碍胸部的视野。引脚应该锋利以顺利地穿刺蛹(图3A)

      图3.隔离IFMs 离体 实时影像。 A.固定蛹。首先切腹腹部(1)。然后,在两侧切割腹侧角质层(2和3),最后水平切开以去除腹角角质层(4)。 (B-B')在通过透射(B)和荧光(B')光去除脂肪和其他不相关的组织之前打开蛹。注意,在这个阶段不能看到IFM,因为它们轻轻地粘附到背角角质层并被周围组织遮蔽。 (C-C')在通过透射和荧光(C)或单独荧光(C')观察到的其它组织清除后的蛹。 IFM圈出。 (D-D')通过透射和荧光(D)或单独荧光灯(D')观察的隔离的IFM对。刻度棒=200μm(A),50μm(D)。

    5. 在双目镜下,用显微切割剪刀剪掉腹部角质层的后端。
    6. 用剪刀,小心地向上滑动并切割两侧的角质层的侧面。穿过睾丸下方的切口,允许移除并丢弃角质层的腹侧部分。
    7. 在荧光双目镜下,放置蓝光滤光片可视化GFP。用镊子,轻轻地去除脂肪体,肠和其他组织,避免与背部角质层和附着的组织接触。在这些阶段,IFM应该轻轻地粘附到背角(图3B和3B')。避免在此阶段清除气管。
    8. 根据使用的特定荧光线,可能还有其他表达GFP的组织。在这些阶段,除了GFP荧光之外,IFM应该在形态学上被识别为6个细长的纤维,轻度地附着在胸背部前部的背侧角质层的两侧。没有GFP荧光的可视化IFM更具挑战性。
    9. 用镊子轻轻清除剩余的脂肪和气管,以发现和暴露IFM。在某些情况下,IFM轻轻连接到气管,必须用镊子分开(图3C和3C),有关其他提示,请参阅“注意”部分)。
    10. 一旦IFM暴露出来,夹住IFM下的镊子将其从角质层上分离出来。
    11. 通过轻轻地将它们用镊子的一侧推进到解剖蛹之外的介质的区域(图3D和3D')来隔离IFM。
    12. 转移200μl施耐德培养基+ 10%FBS,充氧至ibidi板中的Matrigel涂层孔。
    13. 用微量移液管,带有干净介质的湿式200μl微量吸头。在硅板中,将少量液体提取到尖端,并用微量吸管仍然按压,将隔离的IFM提升到50μl液体中。
    14. 轻轻地将IFM和液体转移到中等填充的井中。 IFM应该沉入井表面。
    15. 在镊子的侧面,轻轻按压,压平和定向IFM,以便在ibidi滑块上进行成像。
      注意:任选地,可以将化学抑制剂(例如LatrunculinA)或染料(例如,SiR微管蛋白)加入到相关浓度的培养基中。
    16. 立即使用共聚焦显微镜系统进行成像(例如,蔡司780)。孤立的IFM可以远离成像长达1.5小时。延时电影采用40x水浸物镜,适用于厚生物样品。整个肌肉约35微米厚。使用Zen软件(ZEISS),经常对采集的z-叠层或Z-叠层子集的最大强度投影进行分析。以下采集设置允许以每堆约1.5分钟的时间分辨率对IFM的大部分深度进行成像:33片1.1μm厚度,512 x 512分辨率,线平均为4,2x数字变焦。为了更好的时间分辨率,减少平均值和/或切片数量。

数据分析

一般来说,随着时间的推移可视化可以通过观察所获取的数据的时间延迟来提供对动态行为的洞察。使用Zen软件(ZEISS)创建单片的延时电影或数据的最大强度投影。例如,从表达Lifeact-GFP的切割的IFM的离体培养物到标记肌动蛋白纤维(视频1)的单个光学切片的延时电影显示出从它们的边缘发出的基于肌动蛋白的突起的成肌细胞,其显现为彼此纠缠在一起随着时间的推移,成肌细胞令人惊奇地不动。

Video 1. Ex-vivo IFM expressing mef2 > Lifeact-GFP. Movie shows a single slice of outer surface of isolated IFM, displaying the many myoblasts associated to the myotube.

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具体特征的长度,频率,等等的量化也可以在该系统中执行。例如,Segal等人2016测量了表达膜标记Gap-GFP(Ritzenthaler等人,2000)的IFM中丝状伪足的长度。在这种情况下,随着时间的推移,动力学并不是必需的,因此获取了单个时间点的z-stack(图4)。使用Zen Black软件,选择z片的一个子集来包括肌管的丝状伪足,但不包括来自相邻肌管的背景。然后,进行切片子集的最大强度投影(MIP)。选择感兴趣的区域并进行裁剪(图4,红色框),并调整亮度以突出丝状伪足。将裁剪的原始数据转移到Zen Blue软件,以测量丝状伪足长度。将丝状伪足长度的输出转移到Excel进行分析。在R软件中进行实验组之间的丝状伪足长度差异的统计分析,但可以在任何统计软件上进行。


图4.如Segal等人,2016年的丝状伪足长度分析。详见文本。

笔记

  1. 背部肌肉(DLMs)和背上腹肌(DVM),两种类型的IFM仍然存在。 DLM可以被区分为两个肌肉组中较大者。以前的工作专门对DLM进行成像(Segal等人,2016)。
  2. 在开发过程中,DLM从三根纤维分裂成6根纤维,在16-18 h APF之间。因此,如果在此时间范围内解剖,IFM可能处于分裂中。
  3. 理想情况下,完整的DLM将包括3或6根纤维(见上文)和周围的成肌细胞。然而,单独的肌纤维在解剖过程中并不罕见。一般来说,成肌细胞仍然保留与剩余的肌纤维相关联,并且只要个体肌纤维保持完整,这些样品仍然可以成像。

致谢

这项工作是在以色列雷霍波特魏兹曼科学研究所的本尼·希洛教授和埃亚·谢赫特博士的指导下进行的。这项工作得到以色列科学基金会的资助。本协议的部分内容来自(Weitkunat和Schnorrer,2014)。该方案用于(Segal等人,2016)。

参考

  1. Abmayr,SM,Zhuang,S.and Geisbrecht,ER(2008)。  果蝇中的成肌细胞融合。方法Mol Biol 475:75-97。
  2. Chen,EH和Olson,EN(2004)。 Towards在果蝇中成肌细胞融合的分子途径。 Trends Cell Biol 14(8):452-460。
  3. Dahanyasi,N.,Segal,D.,Shimoni,E.,Shinder,V.,Shilo,BZ,VijayRaghavan,K.and Schejter,ED(2015)。  表面接合和多个细胞接触促进飞行肌肉中的成肌细胞融合 211(1):191-203。
  4. Dutta,D.和VaijayRaghavan,K.(2006)。变态和成年肌肉组织的形成。在In:Muscle Development in< drosophila 中。 Springer 125-142。
  5. Gildor,B.,Schejter,ED和Shilo,BZ(2012)。  双向Notch激活抑制成虫成虫成肌细胞中的融合能力。发育 139(21):4040-4050。
  6. Mukherjee,P.,Gildor,B.,Shilo,BZ,VijayRaghavan,K.和Schejter,ED(2011)。< a class =“ke-insertfile”href =“http://www.ncbi.nlm。 nih.gov/pubmed/21558381“target =”_ blank“>成年果蝇发生期间,肌动蛋白成核剂WASp是成肌细胞融合所必需的。 138(11 ):2347-2357。
  7. OAS,ST,Bryantsev,AL和Cripps,RM(2014)。  J Cell Biol 206(7):895-908 。
  8. Richardson,B.,Beckett,K。和Baylies,M.(2008)。可视化果蝇中的新维度 myoblast融合生物学 30(5):423-431。
  9. Ritzenthaler,S.,Suzuki,E.和Chiba,A。(2000)。< a class =“ke-insertfile”href =“http://www.ncbi.nlm.nih.gov/pubmed/11017174”目标=“_ blank”>肌肉细胞突触后丝状伪足与神经运动神经元轴突相互作用。 Nat Neurosci 3(10):1012-1017。
  10. Schnorrer,F.,Schonbauer,C.,Langer,CC,Dietzl,G.,Novatchkova,M.,Schernhuber,K.,Fellner,M.,Azaryan,A.,Radolf,M.,Stark,A.,Keleman ,K. and Dickson,BJ(2010)。  Systematic遗传分析肌肉形态发生和功能在果蝇。自然 464(7286):287-291。
  11. Segal,D.,Dhanyasi,N.,Schejter,ED和Shilo,BZ(2016)。  肌肉细胞的粘附和融合通过丝状伪足促进。 Dev Cell 38(3):291-304。
  12. Shwartz,A.,Dhanyasi,N.,Schejter,ED和Shilo,BZ(2016)。  果蝇 formin Fhos是肌节细丝阵列组件的主要媒介。 Elife 5.
  13. Volk,T。(1999)。单打出>果蝇肌腱细胞:两种不同细胞类型之间的对话。趋势Genet 15(11):448-453。
  14. Weitkunat,M.,Kaya-Copur,A.,Grill,SW and Schnorrer,F。(2014)。张力和抗力附着对于果蝇中的肌纤维生成至关重要。 ):705-716。
  15. Weitkunat,M.和Schnorrer,F.(2014)。研究果蝇指南肌肉生物学 方法 68(1):2-14。
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引用:Segal, D. (2017). Live Imaging of Myogenesis in Indirect Flight Muscles in Drosophila. Bio-protocol 7(13): e2377. DOI: 10.21769/BioProtoc.2377.
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