We thank Lingfeng Tang for establishing an earlier version of the LarvaSPA method; Glenn Swan at Cornell Olin Hall Machine shop for making earlier prototypes of the imaging chamber; Philipp Isermann for constructing metal frames and providing suggestions on making PDMS cuboids; Cornell BRC Imaging facility for access to microscopes (funded by NIH grant S10OD018516); Maria Sapar for critical reading of the manuscript. This work was supported by a Cornell Fellowship awarded to H.J.; a Cornell start-up fund and NIH grants (R01NS099125 and R21OD023824) awarded to C.H. H.J. and C.H. conceived the project and designed the experiments. H.J. conducted the experiments. H.J and C.H. wrote the manuscript.

1. Making the imaging chamber
<ol>
	<li>The metal frame can be constructed from an aluminum block in a typical machine shop. The specifications of the frame are illustrated in Figure 1A.</li>
	<li>To construct the imaging chamber, seal the bottom of the metal frame using a long coverslip (22 mm x 50 mm) and UV glue (Figure 1A). Cure the UV glue using a hand-held UV lamp.</li>
</ol>
2. Making PDMS cuboids
<ol>
	<li>Prepare the mold for PDMS cuboids.
	<ol>
		<li>Attach layers of packaging tape to the inner surface of a rectangular (80 mm x 55 mm) Petri dish or round cell culture plate. Use one layer (0.063 mm thick) for second instar larvae, 2 layers (0.126 mm thick) for early third instar larvae, or three layers (0.189 mm thick) for late third instar larvae (Figure 1B).</li>
		<li>Cut the tape into strips of specific width with a razor blade: 1.5 mm for the 1-layer tape, and 2 mm for 2-layer or 3-layer tape. The width and thickness of the strip will determine the size of the larvae that the final PDMS cuboid can hold. Leave at least a 5 mm space between the two strips. Remove the tape layers covering the space (Figure 1B).</li>
		<li>Remove dust from the inner surface of the plate using sticky tape. The mold is ready for use.</li>
	</ol>
	</li>
	<li>Prepare the PDMS mix.
	<ol>
		<li>Mix 7 g of PDMS base and 0.7 g of curing agent (10:1 ratio) thoroughly in a small container.</li>
		<li>Place the container in a vacuum desiccator for at least 15 min to remove air from the mixture.</li>
		<li>Slowly pour about 5.5 g of PDMS mixture onto the mold to reach a 1&#8211;2 mm thickness (Figure 1B).</li>
		<li>Place the PDMS mixture in the vacuum desiccator again for at least 15 min to remove remaining air bubbles from the mixture. Break the last few bubbles with a pipette tip.</li>
		<li>Cure the PDMS on a flat surface in a heat incubator at 65 &#176;C for 2 h.</li>
		<li>Use a razor blade to loosen the cured PDMS along the edge of the mold and detach it from the mold. Store the PDMS between two pieces of large sticky tape at room temperature.</li>
		<li>For early and late third instar larvae, cut the PDMS into 8 mm x 2 mm x 1 mm cuboids (along the dotted lines in Figure 1B) by positioning the groove created by the tape strip (step 2.1.2) at the center of the long side of the cuboid (Figure 1B,C). For second instar larvae, cut the cuboid to 8 mm x 1 mm x 1 mm.</li>
	</ol>
	</li>
</ol>
3. Mounting larvae for long-term time-lapse imaging
<ol>
	<li>Prepare the top coverslip for mounting.
	<ol>
		<li>Choose six PDMS cuboids with grooves matching the sizes of the larvae. Follow the recommended groove and size of PDMS based on steps 2.1.1, 2.1.2, and 2.2.7.</li>
		<li>Remove dust from the surface of the PDMS with sticky tape.</li>
		<li>Attach four pieces of double-sided tape (12 mm x 5 mm) on a long coverslip (22 mm x 50 mm) for fixing PDMS cuboids later. The spaces between the two pieces of double-sided tape should be the same as the width of the PDMS groove.</li>
		<li>Apply a small drop (~1.2 &#956;L) of UV glue into the groove of each PDMS cuboid and add six small drops of UV glue into the space between the double-sided tape on the coverslip.</li>
	</ol>
	</li>
	<li>Prepare the larvae for mounting.
	<ol>
		<li>Using a pair of forceps, clean the larvae in water to remove food from the body surface.</li>
		<li>Place clean larvae on a small piece of moistened tissue paper in a small (35 mm) Petri dish without a lid. Place the small Petri dish into a large (60 mm) Petri dish containing a piece of dry tissue paper. In a chemical hood, apply 8&#8211;12 drops (160&#8211;240 &#956;L) of isoflurane onto the dry tissue paper using a plastic transferring pipette and close the lid of the large Petri dish.</li>
		<li>Wait 2&#8211;3 min while monitoring the larvae. Take out the larvae from the large Petri dish once their mouth hooks stop moving.</li>
	</ol>
	</li>
	<li>Mount the animals.
	<ol>
		<li>To image structures on the dorsal side of the animal, place the immobilized larvae onto the UV glue between the double-sided tape on the coverslip with the dorsal cuticle facing the coverslip.</li>
		<li>Cover each larva with a PDMS block and fit the trunk of the larva into the groove of the PDMS. Leave the head and the tail of the larva outside the PDMS groove. Avoid blocking the spiracles of the larva by the glue.</li>
		<li>Press down on the ends of the PDMS block onto the double-sided tape without applying force on the groove. Gently pull on the tail of the larva to flatten the cuticle under the PDMS.</li>
		<li>Cure the UV glue for 4 min using a hand-held UV lamp at the high setting (at about 0.07 mW/mm2). 
		CAUTION: Protect eyes with safety glasses while using the UV lamp.</li>
		<li>Flip the coverslip upside down and repeat step 3.3.4.</li>
		<li>Moisten a small piece of lens paper (15 mm x 30 mm) with 20 &#181;L&#8211;30 &#181;L of water. Place the moistened lens paper at the bottom of the imaging chamber (Figure 1A,D).</li>
		<li>Place the coverslip on the chamber so that the larvae are facing the inside of the chamber. Adhere both ends of the coverslip to the metal surface using UV glue (Figure 1A,D). The dorsal side of the larvae is ready for imaging under confocal microscope (Figure 1E).</li>
	</ol>
	</li>
</ol>
4. Imaging
<ol>
	<li>Image larvae with an appropriate microscope. All results shown in this protocol (Figure 2, Figure 3, Video S2, Video S3, and Video S4) were acquired using a confocal system with a 40x (1.30 NA) oil objective.</li>
</ol>
5. Recovery of imaging chamber and PDMS cuboids
<ol>
	<li>After imaging, remove the oil on the top coverslip using a lens paper. Detach the top coverslip from the metal frame by cutting into the space between the coverslip and the metal frame with a razor blade. The imaging chamber is ready for reuse.</li>
	<li>Detach the PDMS cuboids from the top coverslip with forceps. Roll the PDMS cuboids on sticky tape to remove glue residue and dust. The PDMS cuboids are ready for reuse.</li>
</ol>

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The authors declare no competing interests.

Here we describe LarvaSPA, a versatile method of mounting live Drosophila larvae for long-term time-lapse imaging. This method does not require recovering or remounting larvae, enabling uninterrupted imaging. It is therefore ideal for tracking biological processes that take hours to complete, such as dendrite degeneration and regeneration. This method can be also used for imaging intracellular calcium dynamics and subcellular events such as microtubule growth. As the larval body wall is stable during the imaging...

Time-lapse live imaging is a powerful method for studying dynamic cellular processes. The spatial and temporal information provided by time-lapse movies can reveal important details for answering cell biology questions. The Drosophila larva has been a popular in vivo model for investigations using live imaging because its transparent body wall allows for noninvasive imaging of internal structures1,2. In addition, numerous genetic tools are available in Drosophila to fluorescently label anatomical structures and macromolecules3. However, long-term time-lapse imaging of Drosophila larvae is challenging. Unlike stationary early embryos or pupae, Drosophila larvae move constantly, necessitating immobilization for live imaging. Effective ways of immobilizing live Drosophila larvae include mounting in halocarbon oil with chloroform4, anesthetizing using isoflurane or Dichlorvos solution5, and compressing between the coverslip and the microscope slide6. Although some of these methods have been used for microscopy, none of them is effective for long-term live imaging. Other methods were developed for imaging body wall neurons in crawling larvae using conventional confocal microscopy or light-sheet microscopy7,8,9. However, these methods are not ideal for monitoring cellular dynamics due to the movement of the larvae.
New methods have been developed to achieve long-term time-lapse imaging of Drosophila larvae. Using a polydimethylsiloxane (PDMS) &#34;larva chip&#34;, Drosophila larvae can be effectively immobilized through vacuum-generated suction in a specialized microchamber without anesthetization. However, this method does not offer high temporal resolution for cell biology studies and it has strict limitations on animal size10. Another method using an anesthetization device achieved live imaging of Drosophila larvae at multiple time points and has been applied to study neuromuscular junctions11,12,13,14,15,16. However, this method also does not allow for continuous imaging for longer than 30 min and requires using desflurane repeatedly, which can inhibit neural activity and affect the biological process studied17,18. Recently, a new method that combines microfluidic device and cryoanesthesia has been used to immobilize larvae of various sizes for short periods of time (minutes)19. However, this method requires specialized devices such as a cooling system and longer periods of immobilization require repeated cooling of the larvae.
Here we present a versatile method of immobilizing Drosophila larvae that is compatible with uninterrupted time-lapse imaging for longer than 10 h. This method, which we call &#34;Larva Stabilization by Partial Attachment&#34; (LarvaSPA), involves adhering the larval cuticle to a coverslip for imaging in a custom-built imaging chamber. This protocol describes how to make the imaging chamber and how to mount larvae at a variety of developmental stages. In the LarvaSPA method, the desired body segments are affixed to the coverslip using a UV-reactive glue. A PDMS cuboid additionally applies pressure to the larvae, preventing escape. The air and moisture in the imaging chamber ensure the survival of the partially immobilized larvae during imaging. Advantages of LarvaSPA over other techniques include the following: (1) It is the first method that allows for continuous live imaging of intact Drosophila larvae for hours with high temporal and spatial resolution; (2) The method has fewer limitations on larval size; (3) The imaging chamber and PDMS cuboids can be manufactured at a minimal cost and are reusable.
In addition to describing the larval mounting method, we provide several examples of its application for studying dendrite development and dendrite degeneration of Drosophila dendritic arborization (da) neurons.

<table><tbody><tr><td>6061 Aluminum bars</td><td>McMaster-Carr</td><td>9246K421</td><td></td></tr><tr><td>3M double-sided tape</td><td>Ted Pella, Inc.</td><td>16093</td><td></td></tr><tr><td>3M Scotch Packaging tape</td><td>3M</td><td></td><td>1.88&quot;W x 22.2 Yards</td></tr><tr><td>DUMONT #3 Forceps</td><td>Fisher Scientific</td><td>50-241-34</td><td></td></tr><tr><td>Glass coverslip</td><td>Azer Scientific</td><td>1152250</td><td></td></tr><tr><td>Isoflurane</td><td>Midwest Veterinary Supply</td><td>193.33161.3</td><td></td></tr><tr><td>Leica Confocal Microscope</td><td>Leica</td><td>SP8 equipped with a resonant scanner</td><td></td></tr><tr><td>Lens paper</td><td>Berkshire</td><td>LN90.0406.24</td><td></td></tr><tr><td>Petri dishes (medium)</td><td>VWR</td><td>25373-085</td><td></td></tr><tr><td>Petri dishes (small)</td><td>VWR</td><td>10799-192</td><td></td></tr><tr><td>Razor blade</td><td>Ted Pella, Inc.</td><td>121-20</td><td></td></tr><tr><td>Rectangular petri dish</td><td>VWR</td><td>25384-322</td><td></td></tr><tr><td>SYLGARD 184 kit (PBMS kit)</td><td>Electron Microscopy Sciences</td><td>24236-10</td><td></td></tr><tr><td>Transferring pipette</td><td>Thermo Fisher Scientific</td><td>1371126</td><td></td></tr><tr><td>UV glue</td><td>Norland products</td><td>#6106, NOA 61</td><td>Refractive Index 1.56</td></tr><tr><td>UV lamp (Workstar 2003)</td><td>Maxxeon</td><td>MXN02003</td><td></td></tr><tr><td>Vacuum desiccator</td><td>Electron Microscopy Sciences</td><td>71232</td><td></td></tr><tr><td>Wipes</td><td>Kimberly-Clark</td><td>Kimwipes</td><td></td></tr></tbody></table>

larvaspa, a method for mounting drosophila larva for long-term time-lapse imaging

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because the larval body wall and most internal organs are transparent. However, continuous live imaging of intact Drosophila larvae for longer than 30 min has been challenging because it is difficult to noninvasively immobilizeimmobilizing larvae for a long time. Here we present a larval mounting method called LarvaSPA that allows for continuous imaging of live Drosophila larvae with high temporal and spatial resolution for longer than 10 hours. This method involves partially attaching larvae to the coverslip using a UV-reactive glue and additionally restraining larval movement using a polydimethylsiloxane (PDMS) block. This method is compatible with larvae at developmental stages from second instar to wandering third instar. We demonstrate applications of this method in studying dynamic processes of Drosophila somatosensory neurons, including dendrite growth and injury-induced dendrite degeneration. This method can also be applied to study many other cellular processes that happen near the larval body wall.

The larva imaging chamber is constructed by gluing a custom-made metal frame and two coverslips together. The design of the metal frame is specified in Figure 1A. Drosophila larvae inside the chamber are adhered to the top coverslip with the aid of UV glue and PDMS cuboids. The groove on the PDMS cuboid and the double-sided tape the cuboid is attached to create the space to hold the larvae (Figure 1B,C). The PDMS also applies gentle pressure to ...

Watch this Scientific Journal Video about LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging at JoVE.com

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

This protocol describes a method for mounting Drosophila larvae to achieve longer than 10 h of uninterrupted time-lapse imaging in intact live animals. This method can be used to image many biological processes close to the larval body wall.

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because ...

larvaspa-method-for-mounting-drosophila-larva-for-long-term-time

Research

JoVE Journal

Biology

7.4K Views.  Cornell University. This protocol describes a method for mounting Drosophila larvae to achieve longer than 10 h of uninterrupted time-lapse imaging in intact live animals. This method can be used to image many biological processes close to the larval body wall.

Video: LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Developmental Biology

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Neuroscience

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of the nervous system often produce viable larvae and adults that have locomotion defective phenotypes that are difficult to adequately describe with text or completely represent with a single photographic image. Current modes of scientific publishing, however, support the submission of digital video media as supplemental material to accompany a manuscript. Here we describe a simple and widely accessible microscopy technique for acquiring high-quality digital video of both Drosophila larval and adult phenotypes from a lateral perspective. Video of larval and adult locomotion from a side-view is advantageous because it allows the observation and analysis of subtle distinctions and variations in aberrant locomotive behaviors. We have successfully used the technique to visualize and quantify aberrant crawling behaviors in third instar larvae, in addition to adult mutant phenotypes and behaviors including grooming.

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Here we describe a simple and widely accessible microscopy technique to acquire high-quality digital video of Drosophila adult and larval mutant phenotypes from a lateral perspective.

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Imaging Live Brain Explants from Drosophila melanogaster Third Instar Larvae

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a differentiating ganglion mother cell (GMC), which will undergo one additional division to give rise to two neurons or glia. Studies in NBs have uncovered the molecular mechanisms underlying cell polarity, spindle orientation, neural stem cell self-renewal, and differentiation. These asymmetric cell divisions are readily observable via live-cell imaging, making larval NBs ideally suited for investigating the spatiotemporal dynamics of asymmetric cell division in living tissue. When properly dissected and imaged in nutrient-supplemented medium, NBs in explant brains robustly divide for 12-20 h. Previously described methods are technically difficult and may be challenging to those new to the field. Here, a protocol is described for the preparation, dissection, mounting, and imaging of live third-instar larval brain explants using fat body supplements. Potential problems are also discussed, and examples are provided for how this technique can be used.

Author Spotlight: Investigating Asymmetric Cell Division Dynamics: A Protocol for Live-Imaging of Drosophila Larval Brain Explants 

Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological conditions.

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a ...

Extended Live Imaging of Drosophila Pupal Leg Morphogenesis Without Puparium

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Over the past decades, significant progress has been made in understanding the mechanisms of cell fate determination. However, the process by which fate-determined cells form three-dimensional organismal shapes remains unclear. Recent advances in confocal microscopy have facilitated efforts to observe cell dynamics during development through live imaging. The Drosophila melanogaster pupa is ideal for live imaging due to its immobility, transparent pupal cuticle, and the availability of fluorescent reporter lines. A primary challenge for imaging is the puparium, the cuticle surrounding the pupa, which obstructs optical imaging. While previous methods involved either partial or complete removal of the puparium, maintaining pupal viability for extended periods after this procedure has remained challenging. Here, a simple method is presented for days-long live imaging of the Drosophila leg during the pupal stage, involving complete puparium removal. The method includes removing the puparium from a pupa adhered to double-sided tape, followed by assembling a small chamber on a glass-bottom dish to enclose the pupa and a drop of water. This setup is straightforward, reliable, and supports extended pupal survival by preventing desiccation. Long-term live imaging of the Drosophila pupa has significantly contributed to capturing how the adult leg undergoes dramatic three-dimensional structural changes over 2-3 days. These changes include the transient formation of an intriguing structure (the Parthenon-like structure) by epithelial cells, rapid tissue narrowing, joint formation, and bristle elongation. This method is applicable to the observation of various tissues and can potentially be combined with other techniques, such as optical gene induction, to advance the understanding of cell dynamics during the final shape formation of tissues in the pupal stage.

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Here, a protocol is presented for long-term live imaging of the Drosophila leg during the pupal stage, involving complete removal of the puparium. This protocol can also be applied to other tissues.

Over the past decades, significant progress has been made in understanding the mechanisms of cell fate determination. However, the process by which ...

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

During the rapid inflammatory response to tissue damage, cells of the innate immune system are quickly recruited to the injury site. Once at the wound, innate immune cells perform a number of essential functions, such as fighting infection, clearing necrotic debris, and stimulating matrix deposition. In order to fully understand the diverse signaling events that regulate this immune response, it is crucial to observe the complex behaviors of (and interactions that occur between) multiple cell lineages in vivo, and in real-time, with the high spatio-temporal resolution. The optical translucency and the genetic tractability of Drosophila embryos have established Drosophila as an invaluable model to live-image and dissect fundamental aspects of inflammatory cell behavior, including mechanisms of developmental dispersal, clearance of apoptotic corpses&#160;and/or microbial pathogens, and recruitment to wounds. However, more recent work has now demonstrated that employing a much later stage in the Drosophila lifecycle &#8211; the Drosophila pupa &#8211; offers a number of distinct advantages, including improved RNAi efficiency, longer imaging periods, and significantly greater immune cell numbers. Here we describe a protocol for imaging wound repair and the associated inflammatory response at the high spatio-temporal resolution in live Drosophila pupae. To follow the dynamics of both re-epithelialization and inflammation, we use a number of specific in vivo fluorescent markers for both the epithelium and innate immune cells. We also demonstrate the effectiveness of photo-convertible fluorophores, such as Kaede, for following the specific immune cell subsets, to track their behavior as they migrate to, and resolve from, the injury site.

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

Here we present a protocol for live-imaging wound repair and the associated inflammatory response at high spatio-temporal resolution in vivo. This method utilizes the pupal stage of Drosophila development to enable long-term imaging and tracking of specific cell populations over time and is compatible with efficient RNAi-mediated gene inactivation.

During the rapid inflammatory response to tissue damage, cells of the innate immune system are quickly recruited to the injury site. Once at the wound, ...

Immunology and Infection

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Summary

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Chapters in this video

Live-imaging of the Drosophila Pupal Eye

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Live Imaging of Drosophila Larval Neuroblasts

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture