Long-term in vivo imaging of Drosophila larvae


The Drosophila larva has been used to investigate many processes in cell biology, including morphogenesis, physiology and responses to drugs and new therapeutic compounds. Despite its enormous potential as a model system, longer-term live imaging has been technically challenging because of a lack of efficient methods for immobilizing larvae for extended periods. We describe here a simple procedure for anesthetization and uninterrupted long-term in vivo imaging of the epidermis and other larval organs, including gut, imaginal discs, neurons, fat body, tracheae, muscles and hemocytes, for up to 8 h. We also include a procedure for probing cell properties by laser ablation. We provide a survey of the effects of different anesthetics, demonstrating that short exposure to diethyl ether is the most effective for long-term immobilization of larvae. This protocol does not require specific expertise beyond basic Drosophila genetics and husbandry, and confocal microscopy. It enables high-resolution studies of many systemic and subcellular processes in larvae.


The fruit fly Drosophila is one of the most widely used multicellular organisms for the study of general biological processes. Originally, the adult fly was used to work out the rules of chromosomal inheritance; later, the embryo came to be used for the study of the genes that direct development. More recently, Drosophila has been used to study medically relevant problems ranging from wound healing to cancer, neurodegenerative diseases, metabolism and aging1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16. The available genetic techniques enable gene manipulation (both gain- and loss-of-function) with high spatial specificity and temporal control.

Very good reasons exist for adopting fly larvae and pupae as experimental systems. For example, many gene functions are not required, and many important developmental and physiological processes do not occur in the embryo and can therefore not be studied. Processes that can be better or exclusively studied at larval stages in development include growth control, organ innervation, memory formation and behavior, tumorigenesis and others.

Investigation of many scientific questions is aided by live imaging of the processes under study. While all developmental stages—from embryo to larva, pupa and adult fly—have been exploited for experimental analysis, the subject of live imaging in the past has mainly been the embryo. It was one of the early model systems used for live imaging, even before the introduction of GFP, largely because it is translucent and immobile, two important prerequisites for long-term live imaging17,18. The later stages in the Drosophila life cycle are also amenable to live imaging but pose greater technical and physiological challenges19,20,21,22. The pupa has been used for live imaging of a variety of cellular events23,24,25,26,27,28, but dissection is cumbersome and is therefore the stage of choice only in especially justified cases.

The larva is suitable for imaging, but a major obstacle to its use for long-term imaging has been the fact that larvae continuously move and are difficult to immobilize for extended periods. Methods used in the past for imaging larvae relied on immobilization by mechanical means or treatment with anesthetics20,29,30,31,32,33. Drawbacks of these highly useful methods include undesired side effects such as stress through mechanical pressure, low viability of the larvae and the restriction of immobilization to short periods.

We developed a method that overcomes these problems and allows immobilization of larvae over extended (up to 8 h) periods34. We established laser ablation methods to induce defined wounds in the epidermis and to analyze tension in the larval epidermis while avoiding damage to other organs34,35. We have used the method to study aspects of signal transduction during epidermal wound healing34. As we show here, the method also enables imaging of internal organs, including gut, fat body, tracheae, imaginal discs, neurons, muscles and hemocytes, and is suitable for studying many aspects of cellular and subcellular events.

Advantages of Drosophila larvae as a model system

The larva has all of the general advantages of Drosophila as a model system. Transgenic tools such as Gal4 drivers, fluorescently tagged proteins and other markers, which were originally developed and established for studies in embryos, can also be used for larvae. In addition, larvae are resilient to heat shock and UV irradiation, the methods used to induce mosaic clones and to control temperature-sensitive protein function and optogenetic tools34,36,37,38.

In the embryo, many proteins and RNAs are provided maternally and persist throughout embryonic life, which can interfere with early genetic loss-of-function studies39,40,41,42,43. In the larva, most genes can be knocked down very effectively, because most maternally contributed proteins that may mask gene functions in the embryo have been depleted by this stage.

The larva is translucent and can be imaged without dissection. Many tissues are polyploid; as a result of this, the cells are very large. This makes them ideal for imaging and for the study of subcellular processes; they are easily accessible for single-cell manipulations without risking damage to neighboring cells or other organs34.

A range of interesting physiological processes occur during the larval stages of the life cycle. Most importantly, this is the main period of growth of the animal; during this stage the final body size of the adult fly is established. Growth is controlled by conserved signaling pathways, including the nutrient-sensing pathways and most oncogenic pathways44,45,46,47,48,49. Growth can occur by cell proliferation, for example, in imaginal discs and the brain, or through cell growth based on endoreduplication, as in the epidermis or salivary gland50. The patterning of differential growth is controlled by conserved signaling pathways and developmental genes (e.g., Dpp/BMP, Wnt, Hh and EGF pathways), and the larva and its organs present an excellent system with which to analyze stage- and tissue-specific differential organ growth51,52. For these reasons, the Drosophila larva has also become a well-established system for cancer research into tumor initiation, progression and metastasis in different organs53,54,55,56.

Finally, it is easy to treat larvae with drugs such as rapamycin, chloroquine, bafilomycin A1, wormatin, blebbistatin and many others, which can be administered simply by including them in the food34,57,58,59. The larva therefore represents an attractive in vivo model both for fundamental and medically relevant questions.

Development and validation of the protocol for long-term in vivo imaging of Drosophila larvae

On the basis of classic methods originally used to anesthetize adult flies, Galko and colleagues60,61 introduced the use of an anesthetic chamber for brief immobilization of larvae to induce epidermal wounds without subsequent live imaging.

We adapted this anesthetic chamber and developed the protocol described here for immobilizing larvae for extended periods of live imaging34. The protocol we describe here uses diethyl ether for anesthetization. A single, short-term exposure of the larvae to diethyl ether (3–4.5 min, depending on larval stage; see Tables 1 and 2) is sufficient to keep the larvae immobile for up to 8 h.

Table 1 Comparison of different anesthetics for immobilization of larvae
Table 2 Time of exposure to anesthetic for live imaging of larvae in different instar larval stages

We have successfully created epidermal wounds by laser, followed by long-term 4D in vivo confocal imaging (up to 8 h) to document subcellular signaling and the dynamics of wound healing34. We have also made laser cuts and carried out quantitative analysis of cell junctional tension in the epidermis35.

Comparison with other methods

Drosophila larvae have in the past been immobilized for live imaging either by mechanical methods or by anesthetics. One method of mechanical immobilization uses pressure on the larva through water capillary force29. A single larva is pressed between two coverslips with the help of a drop of water, which allows immobilization for short-term (5–10 min) live imaging. Keeping larvae under pressure for >15 min kills them29.

In another method, larvae are immobilized in a microfluidic chip30,62. Two microfluidic devices were designed: one for short-term (up to 1 h) imaging, the other for long-term (up to 12 h) imaging of early third instar larvae. In both devices, a constant vacuum maintains a strong seal between the chip, larva and coverslip interface to immobilize the larva. For long-term immobilization, a pulsed CO2–air mixture is supplied under moderate pressure to the microfluidic chip. Because these methods are based on mechanical pressure and squeezing of the larvae, they may induce acute and chronic stress responses and may affect signaling pathways and gene expression, as well as cause changes in the mechanical tension and shape of the tissues. Supplying CO2 could also interfere with aspects of systemic physiology.

The main anesthetic of choice for larvae has been desflurane20,31,32,33. Anesthesia with desflurane is more efficient than mechanical methods, and larvae survive better20,31. However, one difficulty is that desflurane evaporates rapidly20,31. As soon as it evaporates, the larvae start to move, necessitating repeated treatments for longer observations. The most complex and critical step in this method is thus application of desflurane during imaging.

Fuger et al.20 developed a chamber for continuous desflurane exposure during imaging. The chamber is composed of two complex parts, a vaporizer/anesthesia device and an imaging chamber20,63,64. In this approach, a single larva is repeatedly exposed to desflurane for 5- to 10-min periods with recovery intervals of 5 min, with the cycle repeated up to 10 times. Heemskerk et al.31 developed a chamber for desflurane exposure with a simpler design for multiple rounds of anesthetization distributed over several hours (four exposures over 6 h). This method has the advantage that animals can feed during the recovery periods, which can be important in experiments in which feeding is critical, for example, in the studies of Rasse et al.64 on the development of neuromuscular terminals and their synapses. However, each round of anesthetization requires ~15–30 min of preparation, which includes washing the larvae, a short pre-anesthetization to orient the larvae in the chamber, exposure to desflurane, placing the chamber under the microscope, focusing and adjusting the software.

Because of their height, both of the aforementioned desflurane chambers can be used only on inverted microscopes20,31 and were designed for single larvae, which are individually immobilized and imaged. Thus, they are time consuming and demand extended microscope usage, which means they are not ideal for collecting data for statistical analysis and forward or reverse screening.

In a recent study, Cevik et al.33 developed a simpler protocol for immobilizing up to 5 larvae at a time, without the use of manipulation devices, vaporizers or imaging chambers. A comparison of desflurane and chloroform showed that desflurane arrested the heartbeat in the first few minutes, but this was followed by rapid recovery. Chloroform provided more rapid anesthesia but slower recovery. Using this method to anesthetize the larvae with chloroform or desflurane is appropriate for brief, intermittent periods of in vivo imaging.

Finally, an additional advantage of the method we describe here is that up to 30 larvae can be immobilized and mounted for microscopy simultaneously and imaged in parallel via 4D in vivo imaging using a motorized stage with x, y and z coordinates for each larva (e.g., a spinning disk confocal microscope with a Nanopositioning Piezo Z stage). The time resolution of the imaging that can be achieved depends on (i) the amount of time needed for imaging each single larva (which in turn depends on the objective lens and total magnification), the number of different fluorophores to be imaged, the number of z-sections, the laser power and the exposure time; (ii) the speed with which the microscope stage moves from one position to the next; and (iii) the number of larvae (see the ‘Microscope parameters used for optimal resolution and live confocal imaging’ section in the Procedure). For example, if the image acquisition of a z-series (50-µm depth) for each larva requires 10 s, the entire set of 30 larvae is completed within 7–10 min, which means that each larva is 3D-imaged at a 7- to 10-min temporal resolution. We find that after 1–2 h, most of the larvae are still immobile. Eventually, some larvae start to move, twitch, drift out of focus or die, so that after 5 h of imaging (Fig. 1), we were able to record full movies for up to 20 of the 30 larvae.

Fig. 1: Effect of different anesthetics on larval survival.

The graph shows the proportions of L3 larvae that pupate (light gray) and develop into adult flies (dark gray) after washing, 3.5 min of exposure to anesthetic (diethyl ether, ethyl acetate, isoflurane or desflurane) and 5 h of starvation, which corresponds to 5 h of imaging without nutrient. Average numbers of survivors as pupae and adult flies for each condition tested are shown. The experiments were done in triplicate, with 30 heterozygous larvae for each condition. Data are given as mean ± s.d. Transgene genotypes: A58>Src-GFP, DsRed2-Nuc (w1118; +; A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+).

In summary, the choice of method depends on many parameters. If the process under investigation does not require continuous imaging and may be sensitive to prolonged anesthesia or starvation, then intermittent anesthesia with periods of rest and feeding may be preferable. For processes that require uninterrupted documentation, we find that the anesthetic and the method of application described here are especially suitable.

Applications of the method

Long-term live imaging of Drosophila larvae can be used to visualize many organ systems: epidermis, neurons, tracheae, muscle, fat body, gut, hemocytes, imaginal discs, and others. Any subcellular processes for which there are live imaging markers can be studied, for example, calcium waves34, phagocytosis34, endocytosis, ER or Golgi vesicle trafficking, mitochondrial fusion and division, autophagy and cell death.

The same method can be used to obtain single images as a substitute for immunofluorescence studies of fixed specimens, as long as a fluorescent marker for the protein of interest is available. Anesthetizing and imaging 20–30 larvae in parallel takes 30–120 min, whereas immunofluorescent staining of larval organs takes 2–5 d, and it is difficult to handle >20 larvae for each round of staining. In addition, fixation can change the properties or morphology of the targeted cells and tissues.

In addition to its high efficiency, this protocol is fast and cheap, and can easily be adapted to other chemical anesthetics, such as desflurane, isoflurane and ethyl acetate, as will be discussed below. This method requires no special expertise or instrumentation, assuming that anyone wishing to apply it is already proficient in Drosophila genetics and husbandry, as well as in general confocal microscopy.

Limitations of long-term imaging

Some limitations are inherent to long-term imaging of larvae, irrespective of the method of immobilization. During larval stages the animals normally feed constantly and grow rapidly. Long-term imaging induces two types of stress. First, feeding ceases and the larvae are starved. Indeed after 4–6 h of starvation, autophagy can be seen in most tissues65,66,67. As observed previously in fat body and muscles, autophagosomes first appear in the epidermis after 4.5 h of imaging34. Thus, at least in the first 4.5 h, this anesthetization method does not induce autophagy and can be used without concern. Live imaging for >5 h requires careful control and analysis.

The constant illumination during imaging can also cause stress. The laser beam concentrates energy locally to visualize fluorescent proteins, which can lead to phototoxicity and local heat stress. However, these problems apply to any long-term live imaging experiment in any cell type or organism.

Overview of procedure and experimental design

We describe here a protocol for long-term in vivo confocal imaging of Drosophila larvae. Originally developed to study epidermal wound healing34, this improved protocol can be applied to study any cellular or subcellular events in any organ, as outlined in detail in the Procedure.

Before starting the experiment, it is necessary to select or generate optimal lines for live imaging of the process to be studied (Step 1). The growth of the larvae should be synchronized (Steps 2–3), especially if several are to be imaged in parallel. We provide instructions for the construction of a larval cage and anesthetic chamber (Steps 4–11) and the anesthetization of larvae (Steps 12–22). We describe the microscope parameters for optimal long-term in vivo imaging of epidermis (Step 23A) and internal organs in the larva (Step 23B) and protocols for multi- and single-cell laser ablation for wound healing studies (Step 24A) and laser cutting of cell boundaries to measure junctional tension (Step 24B). Survival of the animals should be monitored so that unhealthy individuals can be excluded from evaluation and quantitative measurements (Steps 25–27). Finally, we describe the import and processing of images (Step 28), and analysis and quantification of wound closure and junctional tension (Steps 29–30) (Fig. 2).

Fig. 2: Outline of the experimental procedure.

Experimental steps (magenta) and the anticipated duration (green) required for each step are summarized in this image.

We also describe how the same equipment and chamber can be used to anesthetize the larvae with anesthetics other than diethyl ether and compare the effects of desflurane, isoflurane, ethyl acetate and diethyl ether on immobilization and survival of the larvae (Tables 1 and 2). We provide a list of Gal4 drivers (Table 3), markers for visualizing subcellular compartments and organelles (Table 4), and useful GFP protein traps, which we tested and found most useful for live imaging in larvae (Table 4).

Table 3 Gal4 lines for imaging various organs in Drosophila larvae
Table 4 Fly stocks for imaging subcellular compartments in Drosophila larvae

Specific considerations for individual sections of the procedure are detailed below.

Selection and generation of lines for live imaging (Step 1)

The experimental expression of genes with the help of the Gal4/UAS system is a common and efficient method of marking tissues or cells of interest and manipulating gene function in Drosophila68.

We recommend testing Gal4 and UAS lines for their specificity and efficiency as part of the design of the experiment, especially if they will be used for gain- and loss-of-function studies. We tested a range of Gal4 driver lines for their tissue specificity (Table 3). None directed expression in only one larval tissue. The selected Gal4 drivers, fluorescent reporter constructs and GFP-traps we used are listed in Tables 3 and 4.

For precise visualization and analysis, the expression of fluorescent markers should ideally be restricted to the tissue of interest so that the surrounding tissues are free of fluorescent proteins. Fluorescent signals from underlying organs, in particular fat body, make it difficult to detect the signal in the tissue of interest.

All red fluorescent markers (mCherry, RFP, Red2, Tomato) that we have tested in the epidermis and other organs, regardless of which proteins they tagged, showed cytosolic aggregates that were seen as very bright spots. These do not necessarily interfere with detecting the fusion protein proper (e.g., when visualizing actomyosin cables or FOXO shuttling)34 but may influence aspects of quantification, especially if small organelles or cytoplasmic vesicles are to be counted. Therefore, quantitative and qualitative analysis with mCherry, RFP, Red2, Tomato, and so on should be performed very carefully and critically.

To express constructs in the epidermis, four Gal4 drivers can be used: e22c-Gal4, A58-Gal4, 69B-Gal4 and da-Gal4 (refs. 61,68,69,70). The best marker constructs for cell outlines and membranes are UAS-Src-GFP (coding for a myristoylated GFP fusion protein) and UAS-mCD8-GFP (coding for a transmembrane GFP fusion protein). They can be imaged for long periods without much bleaching.

The Galko lab established Drosophila larvae for epidermal wound healing experiments and developed a number of useful tools for examining the larval epidermis61,71. To image epidermal wound healing in larvae, we took advantage of the existing tools and used two lines, carrying either A58-Gal4 or e22c-Gal4; each is recombined with UAS-Src-GFP and UAS-DsRed2 (refs. 61,70):

w1118; e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/CyO; + flies and

w1118; +; A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/TM6B flies.

Both drivers, e22c-Gal4 and A58-Gal4, are expressed in the epidermis. e22c-Gal4 is expressed from embryonic stages 10–11 onward and A58-Gal4 from the first larval instar stage onward61,70. In both lines, the plasma membrane is marked by Src-GFP and the nuclei by DsRed2-Nuc34,61.

In the experiments to measure the recoil velocity of the cell cortex after laser cutting, we visualized cell outlines using a transgenic line in which GFP is inserted into the endogenous DE-Cad gene (Drosophila E-cadherin, the single-pass transmembrane protein of adherens junctions) to create a GFP fusion protein (w1118, y; endo:DEcad-GFP; +)35,72. We obtained best results with lines in which DE-Cad is expressed under its own promoter rather than a tubulin or ubiquitin promoter. The endo:DE-Cad-GFP chromosome is homozygous viable and causes no cellular or developmental abnormalities. The transgenic proteins are sufficiently bright to be imaged easily even when the animal is heterozygous for the chromosome.

Staging and synchronizing the larval growth (Steps 2 & 3)

If several larvae are to be imaged simultaneously or mutants are to be compared with control larvae, larval growth must be synchronized to obtain larvae that are at the same developmental stage. This is not only helpful for the experimental design, it is necessary because different stages require different durations of exposure to the anesthetic (Table 2).

Construction of larval cage and anesthetic chamber (Steps 4–10)

An overview of all materials required for this experiment is shown in Fig. 3, and the experimental setup required to anesthetize the larvae and mount them for confocal live imaging is shown in Figs. 46. We used the design from the Galko lab60 for the larval cage (Fig. 4) and anesthetic chamber (Fig. 5), which are easy, cheap and fast to construct.

Fig. 3: Overview of equipment required to anesthetize larvae.

(1) Large plastic pipette; (2) diethyl ether; (3) glass Coplin staining jar with polypropylene screw cap; (4) culture dish (large); (5) culture dish (small); (6) culture dish for imaging with one compartment; (7) culture dish for imaging with four compartments; (8) tap water; (9) empty food vial in which to raise the larvae after imaging; (10) 10-ml glass beaker; (11) black Nylon sheet; (12) self-made larval cage with lid (see Fig. 4); (13) paper napkin; (14) timer; (15) two paintbrushes with real hair, size 1 and size 2/0; (16) spatula; (17) cotton wool; (18) filter forceps; (19) Nylon mesh, 300- to 400-µm pore size; (20) 1.5-ml microcentrifuge tube; (21) Bunsen burner. The numbers in this image correspond to the numbers in the ‘Equipment’ section (for details see the ‘Equipment’ section).

Fig. 4: Construction of the larval cage.

ad, Photos show how to make a cage used for anesthetization of larvae. For details, see Step 4.

Fig. 5: Construction of anesthetization chamber.

af, These photos show the steps required to construct an anesthetization chamber used for immobilization of larvae. For details, see Steps 5–10.

Fig. 6: Anesthetization and mounting the larvae for live imaging.

Sequence of steps to anesthetize and mount the larvae for live imaging. ac, Sort (a), wash (b) and dry (c) the larvae. d, Place larvae into the larval cage. e, Place the larval cage inside the glass beaker. f, Close the screw cap of the anesthetic chamber and expose the larvae to diethyl ether for 3–3.5 min (see Table 2). g,h, Transfer the anesthetized larvae into an imaging dish (g), and sort and orient (h) them under the stereo microscope. i, Now the larvae are ready for imaging. Place the imaging dish under the microscope. For details of the procedure see Steps 11–22.

Anesthetization of larvae (Steps 11–22)

The anesthetics we have tested (diethyl ether, desflurane, isoflurane and ethyl acetate) are liquids and evaporate when placed into the chamber. Because we found that diethyl ether provides the longest anesthetizing effect, we used it in most of our experiments. The larvae remained immobile for 7–8 h if they were exposed for 3–4.5 min to diethyl ether in the anesthetizing chamber (Tables 1 and 2).

To examine the effects on viability of the experimental procedure itself, and of immobilization with four different anesthetic, we scored the further development and survival of treated individuals (Fig. 1). As a baseline control, early third instar larvae were washed and starved for 5 h in an imaging dish without water or nutrients. Experimental animals were either washed and immobilized with the different anesthetics (diethyl ether, ethyl acetate, isoflurane or desflurane) for 3.5 min or subjected to this treatment and starved as well. Each experiment was done in triplicate, with 30 heterozygous A58>Src-GFP, DsRed2-Nuc (w1118; +; A58-Gal4, Src-GFP, DsRed2-Nuc/+) larvae in each sample. After washing and sorting only, 84% of the larvae survived to adult stages. If, in addition, they were starved for 5 h, 80% of the larvae survived until pupariation and 71% survived until eclosion of the adult. These survival rates were not affected further by anesthetization with diethyl ether. After washing and exposure to diethyl ether for 3.5 min without subsequent starvation, 78% of the larvae survived until pupariation and 67% eclosed as adult flies. With the combination of all manipulations—washing, sorting, diethyl ether exposure for 3.5 min and 5 h starvation—64% of the larvae survived until the pupal stage and 60% survived until eclosion as adult flies.

We also tested the effects of other anesthetics on viability, both with and without long-term starvation (Fig. 1). Desflurane exposure resulted in slightly better survival rates than diethyl ether, with or without starvation. Ethyl acetate exposure performed equally to and isoflurane exposure performed worse than exposure to diethyl ether. Of all the anesthetics, we found isoflurane to be the most toxic to Drosophila larvae.

Unlike diethyl ether, 3.5 min of exposure to desflurane, isoflurane or ethyl acetate is not sufficient to anesthetize larvae for uninterrupted long-term live imaging (Tables 1 and 2). To immobilize them for >10–20 min, larvae must be exposed for 8 min to desflurane, isoflurane or ethyl acetate. We have recorded the maximal imaging times during which at least 25–10% of the larvae remained immobile (Table 1). With the same exposure time (3.5 min), diethyl ether provided the longest (7–8 h) and desflurane the shortest (10–20 min) narcotizing effect (Table 2).

The method of choice for immobilizing larvae for any particular application will depend on a range of parameters. For example, it is possible that diethyl ether and other anesthetics may affect physiological or behavioral phenomena that we are not aware of and that our assays do not detect. If intermittent imaging over long periods is a sufficient readout for the effects of an experimental manipulation, then desflurane, with its documented less negative impact on viability, may be the preferred anesthetic. For uninterrupted long-term imaging of the larvae, diethyl ether is a particularly suitable anesthetic.

Microscope setup for optimal long-term in vivo imaging of epidermis and internal organs (Step 23)

Before anesthetizing larvae and starting an experiment, it is important that the microscope be completely set up, with all parameters entered and ready for imaging.

For long- and short-term live image acquisition of epidermis and other organs, a spinning disk confocal microscope is best suited to fast imaging and minimal photobleaching. For any given application, the total exposure time, laser power, frequency of time intervals, objective, number of sections per z stack, and number of fluorescence channels acquired must be optimized empirically to achieve the desired balance between speed of imaging, image quality and photobleaching. The parameters used for long-term live imaging of larval epidermis in our own experiments are described in the Procedure. To trace fast subcellular processes within the epidermis (e.g., calcium flashes or vesicle trafficking), we used faster laser scanning (Procedure).

Live imaging of internal organs in larvae is not trivial. Although the anesthetic stops somatic muscle movement, the peristaltic gut movement and heartbeat continue, and their movements are indirectly transmitted to other internal organs. However, with high-speed imaging at a rate that is considerably faster than the speed of internal organ movements, it is possible to image imaginal discs, fat body, gut, muscles, tracheae, neurons, and hemocytes (Fig. 7, Supplementary Videos 111). The microscope settings for imaging internal organs are described in the Procedure.

Fig. 7: Live imaging of inner organs and subcellular organelles.

a,b, Mitochondria (green) and nuclei (magenta) in the dorsal epidermis of larvae expressing A58>mito-GFP; DsRed2-Nuc (a); higher magnification of epidermal mitochondria during wound healing (b), corresponds to Supplementary Video 1. c,d, ER (green) in the dorso-lateral epidermis of larvae, e22c>; RFP-KDEL (c); higher magnification (d) of area marked in c corresponds to Supplementary Video 2. e,f, Fat body (green), c7>mCD8-GFP (e); higher magnification (f). e and f corresponds to Supplementary Video 3. g,h, Trachea (green) and nuclei (magenta), e22c>Src-GFP,DsRed2-Nuc (g); trachea (green), btl>GFP (h). g and h corresponds to Supplementary Video 4. i,k, Entire wing imaginal disc (green, white arrow), esg>GFP (i); wing pouch region (green), nuclei (magenta), Pdm2>mCD8-GFP; DsRed2-Nuc (j); posterior compartment of wing imaginal disc (gray), hh>mCD8-GFP (k), i and k correspond to Supplementary Video 5. l,m, Peripheral neurons (green) with nucleus (magenta), Nub>mCD8-GFP; DsRed2-Nuc (l); peripheral neurons (green), elav>mCD8-GFP (m), corresponds to Supplementary Video 6. n,o, Midgut enterocytes (green) (n); midgut interstitial cells (green) (o), NP1>GFP-Atg8a; foxo (n,o), high levels of FOXO in enterocytes and interstitial cells induce autophagy (green dots represent autophagosomes (arrows)). n and o correspond to Supplementary Videos 8 and 9, respectively. p,q, Hemocytes (magenta), Hml>dsRed (p); and hemocytes (green), Jupiter-GFP trap (q). p and q correspond to Supplementary Video 10. r,s, Muscle (green), MHC>GFP-Atg8a; foxo (r), overexpression of FOXO in muscle induces autophagy (green dots represent autophagosomes (arrows)); muscle (green), Nrg-GFP trap (s). r and s correspond to Supplementary Video 11. a,b,g,h,m,p,q, Projections of the z-stacks from a time-lapse series. Each frame is a merge of 57 (a,b) or 21 (g,h,m,p,q) planes spaced 0.28 μm apart. cf,il,n,o,r,s, Single-plane images from time-lapse series of different inner organs and subcellular organelles of L3 larvae. Scale bars, 20 μm (a,c,f,h,l,n,o,q,r,s); 10 µm (b,d); 15 µm (m); 30 µm (g); 50 µm (k,p); 80 µm (e,j); 100 µm (i). Transgene genotypes of larvae: w1118; UAS-mito-HA-GFP/+; A58-Gal4, DsRed2-Nuc/+ (a,b); w1118; e22c-Gal4/+; UASp-RFP.KDEL/+ (c, d); w1118; c7-Gal4/+; UAS-mCD8-GFP/+ (e, f); w1118; e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+; + (g); w1118; btl-Gal4, UAS-GFP; + (h); w1118; esg-Gal4, UAS-GFP/+; + (i); w1118; UAS-mCD8-GFP/+; Pdm2-Gal4/UAS-DsRed2-Nuc (j); w1118; +; hh-Gal4, UAS-mCD8-GFP/+ (k); w1118; Nub-Gal4/UAS-mCD8-GFP/+; UAS-DsRed2-Nuc/+ (l); w1118, elave-Gal4/UAS-mCD8-GFP; UAS-mCD8-GFP/+; UAS-mCD8-GFP/+ (m); w1118; NP1-Gal4/UAS-GFP-Atg8a; UAS-foxo/+ (n,o); w1118; Hml-Gal4/+; UAS-dsRed/+ (p); w1118; +; Jupiter-GFP (q); w1118; MHC-Gal4/UAS-GFP-Atg8a; UAS-foxo/+ (r); and w1118, Ngr-GFP; +; + (s). ER, endoplasmic reticulum.

Multi- and single-cell laser ablation for wound healing studies (Step 24A)

The larval epidermis is an attractive model for the study of many cellular processes. Similar to simple epithelia in mammals, the larval epidermis is a monolayered epithelium with a basal lamina. On the outside it is covered with an apically secreted cuticle (Fig. 8)34,73. In the case of epidermal wound healing, the absence of epidermal tissue remodeling (rearrangement or morphogenesis) is an additional advantage.

Fig. 8: Epidermis of Drosophila larva.

a, Schematic diagram illustrating the structure of the larval epidermis. b,c, Transmission electron microscopic image of cross-section of larval epidermis. c, Magnification of b. ac, Epidermal cell is marked in yellow; secreted cuticle is marked in blue. The larvae used for transmission electron microscopy had the following genotype: w1118; e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+; +. Scale bars, 1 μm. BL, basal lamina; C, cuticle; E, epidermis; Hd, hemidesmosome; M, mitochondria; N, neuron (axon).

For optimal documentation of subcellular events immediately after cell wounding, we recommend the use of the same microscope and lens for laser ablation and imaging. The setup and all parameters and detailed experimental procedure are described in the Procedure.

Laser cutting to measure junctional tension (Step 24B)

To analyze the tension along cell–cell junctions, laser ablations can be performed with the settings described in the Procedure and imaged with the same equipment used for wound healing.

Monitoring the survival of the animals and evaluation of data to obtain quantitative measurements (Steps 25–27)

For reliable comparisons and quantifications of cellular dynamics, all larvae included in the study must have the same fitness. Not all larvae survive the laser wounding and live imaging. It is an advantage to be able to recognize weak larvae early and exclude them from experimentation and further observation. Some weak larvae can be recognized and excluded before the beginning of the experiment; others are visibly damaged during the course of the experiment. The most stringent criterion for quality is survival to adulthood after the experiment.

In some cases, low vitality can be judged by visual inspection. One sign of fitness is the larval heartbeat. Larvae in which the heart is not beating should be excluded from the experiment. Another indication that larvae are not fit is a reduction of intensity or an aggregation of fluorescent markers, or appearance of regular, particulate structures (Fig. 9). Larvae in which the fluorescent markers appear even slightly abnormal should also be excluded (Fig. 9b). Indications that larvae are dying during the experiment include loss of the fluorescent marker (Fig. 9a,h), breakdown of the plasma membrane (Fig. 9a,f,g,h), continuing expansion of the wound and disintegration of tissue (Fig. 9g,h), formation of stress fibers (Fig. 9f), nuclear envelope breakdown (Fig. 9d,h,i), appearance of regular particulate or crystal-like structures (Fig. 9e) and fluorescent markers in inappropriate subcellular locations (Fig. 9b,c).

Fig. 9: Signs of reduced larval vitality.

ai, Indications that larvae are unfit or dying during imaging without laser ablation (ae) or after laser ablation (fi). ai, Reduced intensity of fluorescent markers (a,h; arrows); aggregation of fluorescent protein (b,c; arrows); appearance of regular particulate or crystal-like structures (e; arrows); disintegration of tissue, breakdown of the plasma membrane, continuous expansion of the wound, and formation of stress fibers (f,g; arrows); or breakdown of nuclear envelope and appearance of cytoplasmic fluorescent protein in the nucleus or accumulation of fluorescent marker at the nuclear envelop (d,i; arrows). ai, Projections of an image (ae) or a time-lapse series of the wounded epidermis in third instar larvae (fi). Each frame is a merge of 57 planes spaced 0.28 μm apart. Scale bars, 20 μm (ad,fi) and 10 µm (e). Transgene genotypes: e22c>Src-GFP, DsRed2-Nuc (w1118; e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+; +) (a,b,g,h), A58>mCD8-GFP (w1118; +; A58-Gal4, UAS-mCD8-GFP/+) (cf), and e22c>GFP-mCherry-Atg8a (w1118; e22c-Gal4/UASp-GFP-mCherry-Atg8a/+; +) (i). In i, only the green channel is shown.

Importing and processing of images (Step 28)

For visualization and analysis, the imaging data must be processed, converted and imported using appropriate software. We have mostly used the commercially available software Volocity (v.6.3.57). Other commonly used packages are ImageJ/Fiji (National Institutes of Health), an open source image-processing package, and Imaris (Bitplane) a commercially available software widely used for 4D data analysis.

Analysis and quantification of wound closure and junctional tension (Steps 29 & 30)

Depending on available software, the data can be analyzed and quantified as raw data (e.g., in Volocity) or as converted ‘ome.tif’ files (e.g., in Fiji). The details for quantification are described in the Procedure.


Biological materials

  • Drosophila larvae. The Gal4 driver plus UAS fluorescence reporter constructs and GFP-traps we used for imaging are listed in Tables 3 and 4. Further details are provided in the Procedure.



Exposure to any of the below anesthetic gases is a serious safety concern. Avoid breathing mist or vapor. Diethyl ether, desflurane, isoflurane and ethyl acetate must be directly vented out of the room using a certified ventilated fume hood or a biosafety cabinet. Wear protective gloves/protective clothing/eye protection/face protection. Wash face, hands and any exposed skin thoroughly after handling. Store anesthetic gases in a well-ventilated, locked-up place; keep containers tightly closed; and keep away from heat, hot surfaces, sparks, open flames and other ignition sources.

  • Diethyl ether, for analysis (Sigma-Aldrich, Merck, cat. no. 1.00921.1000)


    Diethyl ether is an extremely flammable liquid that can cause severe acute toxicity if swallowed. Inhalation can cause drowsiness and dizziness, and repeated exposure to skin may cause skin dryness or cracking. Keep away from heat, hot surfaces, sparks, open flames and other ignition sources.


    Diethyl ether is the most efficient anesthetic we have tested here for long-term subcellular imaging of larvae.

  • Desflurane (Suprane, 240 ml; McKesson, cat. no. 842158)


    This chemical is hazardous, can cause eye irritation and serious eye damage, and is suspected of damaging fertility or the unborn child and damage to organs through prolonged or repeated exposure.

  • Isoflurane, USP (Forane, 250 ml; NDC no. 10019-360-60)


    Isoflurane is considered hazardous to health and can cause skin irritation, serious eye irritation and reproductive toxicity. A single exposure may cause drowsiness or dizziness.

  • Ethyl acetate (Sigma-Aldrich, EC no. 205-500-4)


    Ethyl acetate is a highly flammable liquid and vapor and is classified as hazardous. It can cause serious eye damage, eye irritation, drowsiness and dizziness (upon a single exposure). Prolonged or repeated contact may dry skin and cause irritation or cracking.



The equipment for anesthetization is shown in Fig. 3. The numbers in the list below correspond to the numbers in the magenta circles in Fig. 3.

  • No. 1: Transfer pipette (polyethylene, 7.1 ml, bulk pack; Sigma-Aldrich, Merck, cat. no. Z350753-100EA)

  • No. 2: Diethyl ether (see ‘Reagents’)

  • No. 3: Glass Coplin staining jar with polypropylene screw cap and 55-ml or 60-ml capacity (Wheaton, cat. no. 900570)

  • No. 4: Plastic Petri dish (diameter 100 mm, polystyrene; Sigma-Aldrich, Merck, cat. no. P5731)

  • No. 5: Plastic Petri dish (diameter 60 mm, polystyrene; Sigma-Aldrich, Merck, cat. no. P5481)

  • No. 6: Culture dish for imaging (CellView cell culture dish, polystyrene, 35/10 mm, glass bottom, one compartment, sterile; Greiner Bio-One, cat. no. 627861)

  • No. 7: Culture dish with four compartments (CellView cell culture dish, polystyrene, 35/10 mm, glass bottom, four compartments, tissue culture, sterile; Greiner Bio-One, cat. no. 627870)

  • No. 8: Tap water at room temperature (21 °C)

  • No. 9: Empty food vial (in which to raise the larva after imaging), one for each larva

  • No. 10: Glass beaker (10 ml; Duran, cat. no. 21 106 08)

  • No. 11: Black Nylon sheet, for better visualization of larvae during washing and sorting (black plastic file folder)

  • No. 12: Self-made larval cage with lid (Figs. 3 and 4)

  • No. 13: Paper napkin (with little fluff)

  • No. 14: Timer

  • No. 15: Two paintbrushes with real hair (not plastic): one for washing the larvae (size 1; Gerstaecker, cat. no. 8-22621) and a finer one for sorting and orienting them (size 2/0; Gerstaecker, cat. no. 8-69656)

  • No. 16: Micro-spatula (LaborShop24, cat. no. 3530-3535)

  • No. 17: Cotton wool

  • No. 18: Filter forceps, blunt ends, stainless steel (Sigma-Aldrich, Merck, cat. no. XX6200006P)

  • No. 19: Nylon mesh (300- to 400-µm pore size, e.g., Nylon filtration tissue (sifting fabric); NITEX, mesh opening = 300 µm; Kisker Biotech, cat. no. 074003)

  • No. 20: Safe-Lock microcentrifuge tubes (1.5 ml; Eppendorf, cat. no. 0030120086)

  • No. 21: Bunsen burner

Additional items

  • Microscope. For all live imaging, we used an inverted spinning disk confocal microscope (Nikon TiE model with Yokogawa, model no. CSU-X1 system) with a Nanopositioning Piezo Z stage control system (NanoScan, model no. OP400, Prior Scientific), Plan-Fluor 40×/1.3 numerical aperture (NA) oil-immersion differential interference contrast (DIC) objective or Plan-Apochromat 60×/1.2 NA water-immersion objective; EMCCD camera (CamLink, model no. C9100-50; 1,000 × 1,000 pixels) controlled by Volocity v.6.3.57, with 488-nm and 561-nm channels with appropriate band-pass filters and dichroic mirrors. The microscope settings were optimized to perform long-term 3D live imaging (1–8 h, with a 2- to 15-min time interval) or fast imaging (with a time interval of some milliseconds) as outlined in this protocol.

  • Ultraviolet laser. All laser ablation and wounding experiments were performed with a 60×/1.2 (for single-cell ablation) or 40×/1.3 (for multi-cell ablation) objective lens on a spinning disk microscope equipped with a 355-nm pulsed ultraviolet laser (Rapp Opto-Electronic, model no. DPSL-355/14; 14-mW mean power, 70 μJ per pulse). Before starting the laser wounding experiments, the laser was calibrated. Parameters used to induce laser wounds or laser cuts on cell–cell junctions are given in the Procedure.

  • Laser power meter (Coherent, LabMax-TOP, 0494F14R).

  • Laser power head sensor and plarizer (Coherent, J-10MT-10KHZ, 0316D14R).



Generation of larvae for wound healing and laser cutting

Timing variable (2–3 d)

  1. 1

    Set up appropriate crosses to obtain the genotypes required for your experiments. Option A details appropriate crosses for setting up models for wound healing; option B provides appropriate crosses to produce larvae for laser ablations at cell interfaces, in which cell outlines are labeled by endo:DE-Cad-GFP.

    1. (A)

      Wound healing

      1. (i)

        Both e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc and A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc chromosomes are kept over a balancer. For imaging of control animals, cross them with wild-type (w1118; +; +) animals:

        P: virgin female w1118; e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/CyO; +


        P: male w1118; +; +

        F1: w1118; e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+; +


        P: virgin female w1118; +; A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/TM6B


        P: male w1118; +; +

        F1: w1118; +; A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+

      2. (ii)

        Carry out further steps of the procedure, imaging with larvae from the F1 generation.

    2. (B)

      Laser cut

      1. (i)

        Set up the following cross with wild-type (w1118; +; +) animals:

        P: virgin female w1118, y; endo:DEcad-GFP; +


        P: male w1118; +; +

        F1: w1118; endo:DE-Cad-GFP/+; +

      2. (ii)

        Carry out further steps of the procedure, imaging the larvae from the F1 generation.

Developmental synchronization of the larvae

Timing 5–6 d


To obtain a sufficient number of animals at the proper developmental stage (early or mid-L3 instar larval stage) the larval growth must be synchronized.

  1. 2

    One day after setting up a cross, let the flies lay eggs in food vials for 30–60 min at 25 °C. Repeat this several times and in parallel in several vials and keep them at 25 °C. Then collect the L3 larvae 5.5–6 d after egg deposition. Not all stocks develop at precisely the same speed, especially if they carry mutations or complex combinations of chromosomes. Thus, this timing must be carefully determined for each stock.

    Critical step

    An alternative, more stringent method for exact synchronization is to follow the protocol of Maimon et al.74: depending on the size of the fly vial, cross 20–30 males with 60–80 virgin females (2–7 d old) in a fly bottle and keep for 3–4 d at 25 °C. For exact synchronization of larvae, let the flies lay eggs for 2–4 h and remove the flies after that. Collect 60–80 first larval instar stage animals 24 h after removal of the flies (i.e., ~24 h after egg laying (AEL), with a 2-h interval for egg deposition), transfer to new food vials and place in an incubator at 25 °C. Collect 25–30 larvae between 72 and 85 h AEL for live imaging.

  2. 3

    Maintain all fly stocks, crosses and larvae at 25 °C under a 12:12 h light/dark cycle at a constant 65% humidity on standard fly food before commencing Step 11.

Construction of the larval cage and anesthetic chamber

Timing ~15 min

  1. 4

    Construction of larval cage with lid. To make a larval cage (Fig. 4), heat the micro-spatula using a Bunsen burner (Fig. 4a) and cut a 1.5-ml microcentrifuge tube 5 mm from the top (Fig. 4b). Carefully melt the cut surface of the top of the tube with the cap and immediately glue it to a Nylon mesh with a 300- to 400-µm pore size (Fig. 4c). Wait 5 min and then cut off the overhanging parts of the Nylon mesh with scissors (Fig. 4d).

  2. 5

    Construction of anesthetic chamber. To make an anesthetic chamber (Fig. 5c), tightly plug the glass Coplin jar with cotton wool (Fig. 5a). To minimize the amount of anesthetic to be used, press the cotton wool very tightly. The maximum volume possible in the Coplin jar should be occupied by compacted cotton wool (Fig. 5a,b), leaving only enough space for a 10-ml glass beaker (Fig. 5f).

  3. 6

    Also plug the 10-ml glass beaker with cotton, but retain enough space to allow the larval cage to be placed into it (Fig. 5b).

    Critical step

    From this step onward, work under a fume hood.

  4. 7

    With the help of a 7.1-ml plastic pipette, fill the Coplin jar with 15–20 ml of diethyl ether (Fig. 5d).


    Exposure to anesthetic gas is a serious safety concern. The diethyl ether or any other anesthetic must be directly vented out of the room using a certified ventilated fume hood or a biosafety cabinet.


    Diethyl ether is sensitive to light and flammable. It can form an explosive vapor, and it should be kept in the dark and cold under a ventilated fume hood. This is also the case for all other anesthetics we have listed in this protocol: desflurane, isoflurane and ethyl acetate.

    Critical step

    The amount of diethyl ether required depends on the free volume remaining in Coplin jar after plugging it with cotton wool and may vary (15–20 ml). There should be enough liquid diethyl ether inside the chamber to saturate the chamber with diethyl ether vapor.

  5. 8

    Place the glass beaker into the Coplin jar (Fig. 5e).

  6. 9

    Also fill the glass beaker with only enough diethyl ether to just soak the cotton wool (in our experience, 2–3 ml; Fig. 5f).


    The diethyl ether inside the Coplin jar should not rise above the middle of the glass beaker, in order to prevent it from flowing into the glass beaker or the larval cage.

    Critical step

    The cotton wool inside the glass beaker should be soaked, but there should be no free liquid.

  7. 10

    Close the Coplin jar tightly.


    Diethyl ether and all other anesthetics listed in this protocol rapidly evaporate, so in each step the anesthetic chamber should be opened for the absolute minimum of time and immediately be shut again.


    The anesthetic chamber (Fig. 5c) must always be closed tightly with its cap and kept under a fume hood, including when no experiments are being performed.


    Diethyl ether is a solvent and can dissolve some synthetic materials. Therefore, direct contact with some types of plastics (e.g., polystyrene) should be avoided.

    Critical step

    Always keep the anesthetic chamber and larval cage in a dry environment and avoid any contact with water. The diethyl ether does not anesthetize reliably if the larvae are damp or wet. The dry air under the fume hood mitigates this.

    Critical step

    For long-term immobilization, we use diethyl ether as anesthetic. For short- and mid-term (<60 min) live imaging, the same anesthetic chamber and larval cage can be used with any of the other anesthetics (Tables 1 and 2 and Fig. 1).

    Pause point

    The anesthetic chamber filled with diethyl ether can be kept for several hours (up to 5 h) at room temperature (~21 °C) under a fume hood. Pauses >5 h are also possible, but because diethyl ether evaporates, the amount must be readjusted before commencing the next step (Step 11).

Anesthetization and mounting of the larvae for imaging

Timing 8–15 min

  1. 11

    To wash the larvae, place both big (100 mm) and small (60 mm) Petri dishes on top of a black Nylon sheet (Fig. 6a). The black Nylon sheet helps the researcher to see and sort the larvae. Fill both Petri dishes with tap water (room temperature). To reduce any chlorine in the tap water, we recommend keeping the water for some days at room temperature, so that the chlorine evaporates.

  2. 12

    Use a metal spatula to scoop the L3 instar larvae out of the soft food into the big Petri dish (Fig. 6a) and wash them thoroughly by gently moving them in the water with the paintbrush (size number 1) to remove the food from the larvae (Fig. 6b).

  3. 13

    Use the same paintbrush to transfer the larvae to the smaller Petri dish. Further remove leftover food by gently moving them in the water (Fig. 6b,c).

  4. 14

    Collect 20–30 larvae one by one with the same paintbrush and dry them on the paper napkin (Fig. 6c).

    Critical step

    This step is essential and could affect the anesthetization. Because diethyl ether and all other anesthetics we have tested here are hydrophobic, the larvae must be gently but thoroughly dried. The moisture of the environment, including that of the larval cage or larva itself, prevents the diethyl ether from penetrating the outer cuticle or the tracheal system and becoming effective.

    Critical step

    To limit any possible stress, keep the washing and drying steps as short as possible (<5 min).


  5. 15

    Transfer the washed and dried larvae into the cap of the larval cage (Fig. 6d). If necessary, carefully dry the larvae inside the cage with a paper napkin.

  6. 16

    To expose the larvae to diethyl ether, open the anesthetic chamber and use the filter forceps to place the larval cage with the cap facing down (with the larvae inside the cap) inside the glass beaker (Fig. 6e).

    Critical step

    Tightly close the cap of the larval cage to avoid contact of the larvae with the diethyl ether–soaked cotton wool inside the glass beaker. Contact with diethyl ether can induce a cold shock in larvae, causing stress or even death.

  7. 17

    Tightly close the screw cap of the anesthetic chamber (Fig. 6f). Late L2 and early L3 larvae should be exposed to diethyl ether for 3 and 3.5 min, respectively. Middle or late L3 larvae should be exposed to diethyl ether for 4–4.5 min (or 5 min at most) (Table 2). However, these times may have to be adjusted for genetically modified or mutant larvae, especially if the genetic condition affects body size.


    The time of exposure to diethyl ether negatively correlates with survival rates of the larvae. To minimize the lethality, the exposure time must be restricted to the exact time mentioned above and in Table 2.

    Critical step

    The chamber can also be used for desflurane, ethyl acetate or isoflurane. Exposure times for these anesthetics vary and also depend on larval growth (size and larval stage). For exact exposure times, see Table 2.


  8. 18

    After the indicated time, open the anesthetic chamber and remove the larval cage with filter forceps.

  9. 19

    Use the moistened fine paintbrush (size 2/0) to transfer the anesthetized larvae one by one by to a 35/10-mm glass-bottom culture dish for imaging (Fig. 6g).

  10. 20

    With the same fine paintbrush, sort and orient the larvae under the stereomicroscope (Fig. 6g–h). If needed, the larvae can be also sorted for fluorescent markers under a fluorescence stereomicroscope before imaging. If the larvae stick to each other and are not easy to move and orient, moisten the brush with tap water. However, after all larvae are in the right position, carefully remove the leftover water with a paper napkin.

    Critical step

    The larvae should be placed at a distance of 2–3 mm from each other (Fig. 6h). As soon as one larva starts to move or is not completely anesthetized, it should be removed from the culture dish; otherwise, it wakes up the other larvae. If one larva wakes up, some of the other larvae also wake up, so it is important to remove any moving larvae immediately to avoid a domino effect. This also needs to be done during imaging. The effect is not caused by the larva moving and touching other larvae. We do not understand the physiological basis of this mysterious phenomenon, which appears to act through an unknown mechanism that does not require direct contact.


  11. 21

    If you are planning to induce a laser wound in the dorsal epidermis (in Step 24) and record the healing process with an inverted microscope, face the dorsal side of the larvae to the glass bottom of the culture dish (as we did; Fig. 6h). To use the correct orientation for the time-lapse recording, always orient the larvae in the same direction for all experiments, for instance, with the head always pointing to the left side (Fig. 6h,i).


    A heartbeat is an indication of whether the anesthetized larvae are alive. The heartbeat can easily be seen from the ventral or lateral sides of the larva under the stereomicroscope. Larvae without a heartbeat should be removed from the experiment.

    Critical step

    For imaging of different organs, larvae must be oriented and imaged from the appropriate side. Image (i) epidermis, neurons and muscles from the dorsal side; (ii) midgut, wing imaginal discs, peripheral neurons and lateral pentascolopidial (Lch5) organ from the lateral side; and (iii) brain and ventral nerve cord from the ventral side. Tracheae, fat body and hemocytes can be imaged from any orientation.

  12. 22

    If using an upright microscope, there is no need to mount the larvae with mounting medium. Put a coverslip (with a thickness suited to the objective lens) on top of the larvae. Avoid pressing the larvae with the coverslip. Proceed to the next step for imaging (Fig. 6i).


    With an inverted microscope, there is no need to use mounting medium, oil or coverslip. The cuticle of the larvae is made of chitin, a long linear sugar73. As soon as larvae are dry and are not in contact with water or in a humid environment, they stick to the glass. Therefore, there is no need to use adhesive or glues to stick them to the culture dish (Fig. 6h,i).

    Critical step

    Besides preventing adhesion of the cuticle to the glass, water could also wake up the larvae. Contact with water should be avoided until the end of the experiment.

Microscope parameters used for optimal resolution and live confocal imaging

Timing variable (up to 8 hours)

  1. 23

    For all our live imaging, we used an inverted spinning disk confocal microscope as described above (‘Equipment’ section)34,35. For live imaging of wound healing, follow option A; for measurement of junctional tension, follow option B.

    1. (A)

      Live imaging of wound healing


      The following microscope settings are suitable for live imaging of A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc or e22c-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc larvae: 488-nm and 561-nm channels (exposure time, 100–180 ms for both 488-nm and 561-m laser beams); 55–65 z-stacks with a step size of 0.28 µm with 60×/1.2 and 0.24 µm with 40×/1.3 (covering a 15- to 20-µm depth), taken every 2–15 min for 1–8 h. These settings do not induce phototoxicity or strong photobleaching during 8 h of live imaging.

      1. (i)

        Set laser power and exposure time. In our experiments, we used the 488-nm laser at 12–15% power and the 561-nm laser at 2–4%. However, because each laser is different and lasers change during their lifetimes, and different constructs have different fluorescent properties, these values must be individually determined.

        Critical step

        To protect the samples against phototoxicity and photobleaching for long-term imaging, reduce the laser power and increase the exposure time (100–200 ms). Longer exposure time obviously reduces the speed of imaging and is not suitable for imaging with narrow time intervals (<1min).

    2. (B)

      Measurement of junctional tension

      1. (i)

        Set the appropriate microscope settings. The microscope settings we recommend for live imaging and laser ablation of endo:DE-Cad-GFP larvae are 488-nm (10–30% laser power in our case) channel (exposure time, 15–60 ms) with a 60×/1.2 or 40×/1.3 objective lens, single planes taken every 0.5 s for 5–7 min, starting 2 min before ablation and finishing 5 min after ablation.


        Increasing the power of the laser beam increases photobleaching and phototoxicity and is recommended only for short-term live imaging. It is important to first run test experiments to find the optimal parameters for each set of experiments with different fluorescent markers .

        Critical step

        These settings (from Step 23B(i)) are also suitable for monitoring fast cellular and subcellular events, for example, vesicle trafficking (Supplementary Video 2) or calcium flashes, which require high-speed imaging at narrow time intervals (some milliseconds).

        Critical step

        These settings (from Step 23B(i)) are also recommended for imaging internal organs over a 5- to 120-min period. For optimal tracking of the organ of interest, reduce the number of z-stacks or choose single-plane imaging. Combine this with a short exposure time (15–60 ms). If necessary, the imaging of different planes can be carried out by manually changing the fine focus during live imaging (Supplementary Videos 3, 7, 8, 9 and 11). This depends on the ability of the software to record and transfer the images simultaneously. This is possible with the Volocity software v.6.3.57 we have used.

        Critical step

        In general, reducing the exposure time and minimizing the number of z-stacks will allow recording at shorter time intervals. However, for better signal-to-noise ratios, higher laser power is desirable.

        Critical step

        For analysis and quantification, it is important to use the same experimental conditions for all genotypes, as well as the same image settings.

Laser ablation

  1. 24

    To undertake multi- or single-cell laser ablation for wound healing studies, follow option A. Follow option B for laser ablation of the cell boundary to measure junctional tension.

    1. (A)

      Multi- and single-cell laser ablation for wound healing studies

      Timing 1 s–10 min

      1. (i)

        Before starting the experiment, correctly calibrate the laser. For a high laser power efficiency, the microscope focus during calibration is extremely important and should be adjusted on the plane nearest to the glass of the imaging dish, for example, on the apical region of the epidermis to be wounded.

      2. (ii)

        Set up a 60×/1.2 (for single-cell ablation) or a 40×/1.3 (for multi-cell ablation) objective lens on a spinning disk microscope equipped with a 355-nm pulsed UV laser (14-mW mean power, 70 μJ per pulse).

      3. (iii)

        Turn on the UV laser and all associated devices. Use an appropriate dichroic mirror to reflect the UV laser into the specimen. For safety reasons, change the shutter to allow bypass of the UV laser. To image events immediately after ablation, use the same microscope and objective lens for laser ablation and imaging.


        Make sure you avoid exposure of your eyes or other body parts to the UV laser beam through the objective lens or ocular. The reflection of light from the laser beam can cause serious damage to the eyes or skin.

      4. (iv)

        Set all laser parameters.

      5. (v)

        Find the dorsal midline of abdominal segment A3 or A4 (or any other region of interest).

      6. (vi)

        Focus on the epidermal cells and make a single image to record the pre-wounding situation. Pause the image acquisition.

      7. (vii)

        Induce a single-cell ablation by focusing on the nucleus in the target cell and marking three spots (diameter for each spot is 6 pixels (px)) on the nucleus with the software of the laser (Fig. 10, Supplementary Video 12).

        Fig. 10: Single- and multi-cell laser wounding in the larval epidermis.

        Time-lapse series of single-cell and multi-cell wound healing in L3 larvae expressing Src-GFP (green) and DsRed2-Nuc (magenta) to mark the cell membrane and nuclei in the epidermis: A58>Src-GFP, DsRed2-Nuc (w1118; +; A58-Gal4, UAS-Src-GFP, DsRed2-Nuc/+). z-projections of a time-lapse series in early L3 larvae. Each frame is a merge of 57 planes spaced 0.28 μm apart. Scale bars, 20 μm. Images correspond to Supplementary Video 12.

        Critical step

        The spot size of the UV laser beam can vary from 1 to 10 px, depending on cell size. For cells of 20- to 30-μm diameter, adjust the UV spot to a 1- to 2-px diameter; for bigger cells (40- to 80-µm diameter) adjust the spot to a 3- to 6-px diameter.

      8. (viii)

        To deliver a laser pulse, click the appropriate icon to release the laser power and shoot the cells with the UV laser beam with 1 pulse/μm laser power of ~0.30 μJ energy. The pulse energy is measured on the objective lens using a laser power meter (half-wave plate) and a laser power head sensor and polarizer form 50 nJ to 12 µJ. This should generate a damaged area of 40- to 60-μm diameter (800–2,500 µm2 area).

        Critical step

        We performed laser ablation during time-lapse imaging without pausing the image acquisition. Alternatively, one can also pause imaging, perform the laser ablation and then continue imaging.

        Critical step

        The highest power of laser beam efficiency is on the focal plane where the beam was calibrated. Therefore, to ablate the cell(s), the correct focal plane must be selected. If laser wounding is unsuccessful, first change the focal plane by changing the fine focus and repeat the laser treatment. If this does not help, gradually increase the laser power and try again until you see an effect. Alternatively, calibrate the laser again or contact the provider to help you for the correct calibration, because this is not a trivial problem.

        Critical step

        If the laser target site is too close to a neighboring cell, damage to the neighboring cell membrane can occur, with deleterious effects on and possibly death of the neighboring cells. If the target site in the cell to be killed is not at the correct level or position, and only cytoplasm is damaged, rather than the nucleus, this may not reliably lead to cell death; sometimes the cells are able to heal themselves, in a process often associated with accumulation and contraction of actomyosin. By contrast, laser ablation of the nucleus always immediately leads to cell death and a response from the neighboring cells, even without damage to the neighboring cells.


      9. (ix)

        For multi-cell ablation, having delivered one laser pulse to a spot of 6-px diameter at the nucleus of each cell to be killed, also deliver a laser pulse to each of the common junctions (Fig. 10, Supplementary Video 12). A multi-cell laser ablation (3–8 cells) can generate a wound of 80- to 120-μm diameter (3,000–8,500 µm2 area).


        The pulse and power of the UV laser beam are the same for both single- and multi-cell ablation (1 pulse/μm, ~0.30 μJ, energy measured on the objective lens). The only difference is the number of positions shot by the laser. For single-cell ablation, three partially overlapping spots are used; for a multi-cell ablation, for example, ablation of three cells, seven different spots must be marked to induce a wound. In all cases, the diameter of the spots was 6 px.

        Critical step

        With a spinning disk wide confocal microscope, larger wounds can be induced because the field of view is 4 times larger than a standard scanning head.

    2. (B)

      Laser ablation of the cell boundary to measure junctional tension

      Timing 1 s–10 min

      1. (i)

        To perform a laser cut at the plane of the cell–cell junction and to avoid a wound healing response use the following UV laser beam setting: 1 pulse/μm of ~0.25 μJ energy (measured on the objective lens).

      2. (ii)

        Find the dorsal midline of abdominal segment A3, A4 or A5 in the microscope.

      3. (iii)

        Focus on the most apical side of the epidermal cells.

      4. (iv)

        Mark the cell boundary with a single spot (diameter for each spot size is 1 px).

        Critical step

        The recoil occurs within some milliseconds; therefore, the laser cut must be induced during the time-lapse imaging to capture and quantify the immediate response to cutting. To image as quickly as possible, set up the microscope for single-plane imaging and image only one channel (488 nm, green, for endo:DE-cad-GFP).

      5. (v)

        Before UV laser ablation, first record a time-lapse series of 2 min (time interval every 0.5 s).

      6. (vi)

        To perform the laser cut, press the appropriate icon to release the laser power (1 pulse/μm of ~0.25 μJ energy, measured on the objective lens).

        Critical step

        If the laser cut is unsuccessful, change the focal plane or increase the laser power or both.

Monitoring the survival of the animals and quantitative evaluation of data

Timing 2–5 d

  1. 25

    To monitor viability after imaging, transfer each larva individually to an empty fly food vial.

  2. 26

    Record on the vial the same name/ID of the larva that was used for recording the imaging data.

  3. 27

    Allow the larva to recover and develop under normal conditions: 25 °C under a 12:12 h light/dark cycle at constant 65% humidity.

    Critical step

    Analyze only the imaging data (next steps) from those animals that survive to become pupae or adult flies. This helps to distinguish the effects of the experimental variables (through genetic manipulation or drug treatment) from the overall fitness of the animals and to avoid analyzing artifacts.

Import and processing of the imaging data

Timing variable


For converting, processing and importing data, any suitable software can be used: Volocity, Imaris, or Fiji. We use Volocity v.6.3.57, Fiji or Imaris for 4D data analysis


For analysis and quantification of the data by Fiji or Imaris, the data must be converted to ome.tif files before importation into those programs.

  1. 28

    Import the metadata to the software (Volocity, Fiji or Imaris), where the data can be analyzed.

    Critical step

    The imaging parameters described in this protocol are adjusted to each fluorescent dye and optimized for long-term live imaging so that fluorescent signal intensity is maintained until the end of live imaging (8 h) and is suitable for qualitative and quantitative analysis.

    Critical step

    The quality of images taken with the microscope settings described here is in general good enough for quantification and analysis.

    Critical step

    If necessary, the threshold of the fluorescence signals can be post-processed in a linear fashion.

Analysis and quantification of wound closure and junctional tension

Timing variable

  1. 29

    To assess wound healing dynamics of the epidermis, define wound areas as the area left open by the dead cells between the lamellipodia. Measure this area manually using Volocity, Fiji or Imaris34.

    Critical step

    Wound healing dynamics of other organs can also be measured in the same way.

  2. 30

    To assess epidermal cell tension, measure the displacement of the cell vertices after laser ablation of the cell bounds over a time interval of 2–5 s, where the first measurement point is the time of ablation35.

    Critical step

    For measurements in other organs, the time interval for imaging should be adjusted (most likely reduced).


Troubleshooting advice can be found in Table 5.

Table 5 Troubleshooting table


  • Step 1, selection and generation of optimal lines for live imaging: variable (2–3 d)

  • Steps 2 and 3, staging and synchronizing of larval growth: 5–6 d

  • Steps 4–10, construction of larval cage and anesthetic chamber: ~15 min

  • Steps 11–22, anesthetization and mounting of larvae: 8–15 min

  • Step 23, setup of microscope and long-term imaging of epidermis or internal organs: variable (up to 8 h)

  • Step 24A, multi- and single-cell laser ablation for wound healing studies: 1 s–10 min

  • Step 24B, laser cutting of cell boundaries to measure junctional tension: 1 s–10 min

  • Steps 25–27, monitoring the survival of the animals and evaluation of data for quantitative measurements: 2–5 d

  • Step 28, import and processing of images: variable

  • Steps 29 and 30, analysis and quantification of wound closure and junctional tension: variable

Anticipated results

Live imaging of systemic, cellular and subcellular processes with high spatiotemporal resolution can improve our understanding of those processes. This protocol enables long-term live imaging of larval organs, subcellular organelles and events in Drosophila larvae (Fig. 7, Fig. 10 and Supplementary Videos 112).

The method has enabled us to detect novel spatiotemporal details of insulin and TOR signaling and their consequences in the wound healing process34. Binding of insulin ligands (Insulin/IGF or Drosophila dILPs) induces the autophosphorylation and activation of the insulin tyrosine kinase receptor (InR/IGFR) and the phosphorylation of insulin substrate (IRS/Chico), which recruits and activates PI3K. Active PI3K catalyzes and increases the level of phosphoinositol (3, 4, 5) triphosphate (PIP3) at the cytosolic site of the plasma membrane, where it serves as a docking site for PIP3­-dependent kinase (PDK) and activates AKT. Activated Akt phosphorylates various target proteins such as FOXO transcription factor and TCS1 and TCS2, inhibitors of the protein kinase TORC1 complex. Once phosphorylated, FOXO accumulates in the cytoplasm and cannot translocate into the nucleus to induce target genes75. To visualize the PI3K activity, we used a common PIP3-sensor, tGPH, and endogenous dFOXO-mCherry34,76. tGPH and dFOXO-mCherry are ubiquitously expressed and distributed throughout the cell. However, during imaging dFOXO-mCherry concentrated in the nucleus. Within 6–9 min after wounding, tGPH became enriched at the plasma membrane and dFOXO-mCherry shuttled out of nucleus in the cells surrounding the wound34 (Fig. 11a and Supplementary Video 13). This indicates an acute activation of insulin receptor signaling (PI3K/FOXO) within minutes during epidermal wound healing.

Fig. 11: Live imaging of insulin/PIP3/FOXO signaling and the effect of reduced insulin receptor signaling on PIP3.

a, Time-lapse images of a single-cell wound in larva expressing tGPH (tubulin:GFP-PH) (green), a PIP3 reporter and endogenous FOXO-mCherry (magenta). PIP3 accumulation (green) and dynamic FOXO (magenta) shuttling between nucleus and cytoplasm in cells directly surrounding the epidermal wound of L3 larvae during wound healing are shown. b, Distribution of tGPH (gray) in the epidermis of control and A58>InRDN larvae. In A58>InRDN larvae, a dominant negative version of the insulin receptor (InRDN) is expressed under the control of the A58-Gal4 driver. Reduced insulin receptor signaling (A58>InRDN) in the epidermis of larvae lowered the PIP3 accumulation (black) at the wound edges and led to wound healing delay. a,b, Projections of a time-lapse series of the wounded epidermis in early third instar larvae. Each frame is a merge of 57 planes spaced 0.28 μm apart. Scale bars, 20 μm. Transgene genotypes: tGPH; foxo-mCherry (w; tGPH; endo-dfoxo-v3-mCherry) (a) and control (w1118; tGPH/+; A58-Gal4/+) and A58>InRDN(w1118; tGPH/+; A58-Gal4/UAS-InRDN) (b). Images correspond to Supplementary Videos 13 and 14.

The epidermis-specific interference with insulin receptor signaling by expression of a dominant-negative version of insulin receptor (InRDN) reduced tGPH accumulation at the plasma membrane and reduced PI3K activity34 (Fig. 11b and Supplementary Video 14). Reduced insulin receptor/PI3K signaling caused a delay of epidermal repair34. We found that the insulin signaling network is needed for the efficient assembly of an actomyosin cable around the wound, and that constitutively active myosin II regulatory light chain suppressed the effects of reduced IIS34.

The most critical steps in this protocol are the steps in which the larvae are exposed to the diethyl ether. Longer exposure times substantially reduce the survival rate of the larvae during or after imaging (Table 5, Step 17). For example, if early L3 instar larvae are exposed to diethyl ether for 6 min instead of 3.5 min (the optimal exposure time for this larval stage), even though they look normal at the beginning of imaging, large numbers of larvae die before the experiment is complete (Fig. 9g and Supplementary Video 15). On the other hand, if the exposure to diethyl ether is too short or the larvae in the cage are too crowded (>30 larvae) or they are not completely dry (Table 5, Step 14), they wake up before the end of the experiment (Supplementary Video 16). Therefore, we recommend using the exact exposure times given in Table 2.

In summary, the immobilization method and long-term in vivo imaging described here, coupled with genetic manipulations, enable the study of many aspects of biology and physiology.

Reporting Summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

All data generated and analyzed during the current study are available from the corresponding authors upon request.


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We are grateful to M. Kakanj for her photographic and graphical support. We thank S. Roth, N. Riddiford, A. Schauss, F. Papagiannouli, V. Böhm and C. Lesch for critical reading of manuscript, comments and helpful discussions. We thank A. Schauss, P. Zentis and C. Jüngst from the CECAD imaging facility in Cologne (University of Cologne, Cluster of Excellence in Ageing Research) for support and the Bloomington, VDRC and DGGR stock centers for fly strains. This work was supported by grants from the European Regional Development Fund and the German state North Rhine-Westphalia (NRW im Ziel 2) to S.A.E. and L.P., a CMMC grant to M.L. and S.A.E., a CECAD grand by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) under Germany’s Excellence Strategy, grant EXC-2030/1-390661388 to M.L. and an EMBL research fellowship to P.K.

Author information

P.K. conceived, designed and developed the long-term live imaging and laser ablation protocol for Drosophila larvae; performed all experiments; and prepared the figures and tables. P.K., M.L. and L.P. analyzed and discussed the data and drafted the manuscript. S.A.E. provided input on wound healing analysis and discussed the data.

Correspondence to Parisa Kakanj or Linda Partridge or Maria Leptin.

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The authors declare no competing interests.

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Peer review information Nature Protocols thanks Greg Macleod and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

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Key references using this protocol

Kakanj, P. et al. Nat. Commun. 7, 12972 (2016): https://doi.org/10.1038/ncomms12972

Beati, H. et al. J. Cell Biol. 217, 1079–1095 (2018): https://doi.org/10.1083/jcb.201610098

Supplementary information

Supplementary Video 1 | Live imaging of mitochondria during epidermal wound healing.

Live imaging of mitochondria (green) dynamics during epidermal wound healing. Single-cell wound in the dorsal epidermis of Drosophila L3 larva expressing mito-GFP (green) and DsRed2-Nuc (magenta) to mark mitochondria and nuclei in the epidermis: A58>mito-GFP; DsRed2-Nuc (w1118; UAS-mito-HA-GFP/+; A58-Gal4, DsRed2-Nuc/+). Each frame is a merge of 57 planes spaced 0.28 μm apart. Scale bar: 20 μm. Corresponds to Fig. 7a-b.

Reporting Summary

Supplementary Video 2 | Live imaging of ER vesicle trafficking.

Live imaging of ER vesicle (green) trafficking between ER and Golgi in the epidermis of Drosophila L3 larva expressing e22c>; RFP-KDEL (w1118; e22c-Gal4/+; UASp-RFP.KDEL/+). KDEL is an ER marker, encodes a sequence to prevent secretion of the protein form ER or retrieval of ER proteins from the Golgi. Each frame is a single plane. Scale bar: 20 μm. Corresponds to Fig. 7c-d.

Supplementary Video 3 | Live imaging of fat body.

Live imaging of fat body (green) in the L3 larvae expressing c7>mCD8-GFP (w1118; c7-Gal4/+; UAS-mCD8-GFP/+). The mCD8-GFP marks the cell membranes. Each frame is a single plane. Scale bars: first movie 80 µm and second movie 20 μm. Corresponds to Fig. 7e-f.

Supplementary Video 4 | Live imaging of trachea.

Live imaging of trachea (green) in the early L3 larvae in two individuals both expressing btl>GFP (w1118; btl-Gal4, UAS-GFP; +). Each frame is a merge of 21 planes spaced 0.28 μm apart. Scale bar: (a,b) 17 μm. Corresponds to Fig. 7g-h.

Supplementary Video 5 | Live imaging of wing imaginal disc.

First movie shows a live imaging through the z-stacks of dorsal compartment of wing imaginal disk (grey) in late L3 larva expressing hh>mCD8-GFP (w1118; +; hh-Gal4, UAS-mCD8-GFP/+). The mCD8-GFP marks the cell membranes. Each frame is a single plane of z-stacks (54 stacks with 1µm space). Scale bar: 50 μm. Second movie shows a short live imaging of entire imaginal disk (green) in early L3 larvae expressing esg>GFP (w1118; esg-Gal4, UAS-GFP/+; +). Each frame is a single plane. Scale bar: 100 μm. Corresponds to Fig. 7i-k.

Supplementary Video 6 | Live imaging of peripheral neurons.

Live imaging of peripheral neuron (black) in the L3 larva expressing mCD8-GFP (black) to mark cell membranes of the neurons: elav>mCD8-GFP (w1118, elave-Gal4/UAS-mCD8-GFP; UAS-mCD8-GFP/+; UAS-mCD8-GFP/+). Each frame is a merge of 21 planes spaced 0.28 μm apart. Scale bar: 15 μm. Corresponds to Fig. 7m.

Supplementary Video 7 | Live imaging of a chordotonal organ.

Live imaging of lateral pentascolopidial (Lch5) organ (green) in L3 larva expressing mCD8-GFP (green) to mark cell membranes: elav>mCD8-GFP (w1118, elave-Gal4/UAS-mCD8-GFP; UAS-mCD8-GFP/+; UAS-mCD8-GFP/+). The Lch5 organ is a particular chordotonal organ, which plays a role by proprioceptive locomotion control. Each frame is a single plane. Scale bar: 8 μm.

Supplementary Video 8 | Live imaging of midgut enterocytes.

Live imaging of midgut enterocytes cells (green) in the L3 larva expressing NP1>GFP-Atg8a; foxo (w1118; NP1-Gal4/UAS-GFP-Atg8a; UAS-foxo/+). Overexpression of FOXO in the midgut enterocytes induces autophagy (green dots are autophagosomes). The fine-focus was changed during the live imaging to image through different planes. Each frame is a single plane. Scale bar: 20 μm. Corresponds to Fig. 7n.

Supplementary Video 9 | Live imaging of midgut interstitial cells.

Live imaging of midgut interstitial cells (green) in the L3 larva expressing NP1>GFP-Atg8a; foxo (w1118; NP1-Gal4/UAS-GFP-Atg8a; UAS-foxo/+). Overexpression of FOXO induces autophagy in the midgut interstitial cells (green dots are autophagosomes). The fine-focus was changed during the live imaging to image through different planes. Each frame is a single plane. Scale bar: 20 μm. Corresponds to Fig. 7o.

Supplementary Video 10 | Live imaging of hemocytes.

Live imaging of haemocytes in the L3 larva expressing Hml>dsRed (magenta, in the first movie) to mark the entire cytoplasm of the haemocytes and Jupiter (green, in the second movie) to mark the microtubules of the haemocytes: Hml>dsRed (w1118; Hml-Gal4/+; UAS-dsRed/+) and Jupiter-GFP trap (w1118; +; Jupiter-GFP). Each frame is a merge of 21 planes spaced 0.28 μm apart. Scale bars: 20 μm. Corresponds to Fig. 7p-q.

Supplementary Video 11 | Live imaging of muscle.

First movie shows live imaging of dorsal muscles (green) in L3 larva expressing MHC>GFP-Atg8a; foxo (w1118; MHC-Gal4/UAS-GFP-Atg8a; UAS-foxo/+), in which high level of FOXO induces autophagy (green dots are autophagosomes). Second movie shoes live imaging of dorsal muscles (green) in L3 larva expressing Nrg-GFP t

Supplementary Video 12 | Single- and multicell laser ablation and wound healing process.

Live imaging of single-cell (first movie) and multi-cell (second movie) laser wounding in the dorsal epidermis of L3 larva expressing Src-GFP (green) and DsRed2-Nuc (magenta) to mark cell membrane and nuclei in the epidermis: A58>Src- GFP,DsRed2-Nuc (w1118; +; A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+). Each frame is a merge of 57 planes spaced 0.28 μm apart. Scale bars: 20 μm. Corresponds to Fig. 10.

Supplementary Video 13 | Dynamic insulin/PIP3/FOXO signaling during wound healing.

Redistribution of PIP3 reporter, tGPH (green) and FOXO (magenta) shuttling after wounding in the cells directly surrounding the wound. Wound healing was performed in the dorsal epidermis of L3 larva expressing tGPH (tubulin:GFP-PH) and foxo-mCherry (w-; tGPH; endo-dfoxo-v3-mCherry). Each frame is a merge of 57 planes spaced 0.28 μm apart. Scale bar: 20 μm. Corresponds to Fig. 11a.

Supplementary Video 14 | Effect of reduced insulin receptor signaling.

Lowering of insulin receptor signalling in the larvae by expressing a dominant negative version of the insulin receptor (InRDN), reduced accumulation of the PIP3-reporter tGPH (back) at the wound edges and slows down the wound healing processes. Transgene genotypes: control (w1118; tGPH/+; A58-Gal4/+) and A58>InRDN (w1118; tGPH/+; A58-Gal4/UAS- InRDN). Each frame is a merge of 57 planes spaced 0.28 μm apart. Scale bar: 20 μm. Corresponds to Fig. 11b.

Supplementary Video 15 | Long exposure to anesthetic increases larval lethality during live imaging.

The early L3 instar larva expressing Src-GFP (green) and DsRed2-Nuc (magenta) was exposed for 6 min instead of 3.5 min to diethyl ether. A single-cell laser wound was made in the dorsal epidermis. The larva died during the experiment. Transgene genotypes of larvae: A58>Src- GFP,DsRed2-Nuc (w1118; +; A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+). Projections of a time-lapse series. Each frame is merged from 57 planes spaced 0.28 μm apart. Scale bars: 20 μm. Corresponds to Fig. 9g.

Supplementary Video 16 | Insufficient immobilization for long-term live imaging.

Example of a larva that did not remain immobile for the entire experiment. The reason for the larva waking up could be that (i) the exposure to diethyl ether was to short or (ii) the larvae in larval cage were too crowed or (iii) they were not completely dried (Table 5, Step 17). A single-cell laser wound was produced in the dorsal epidermis of a larva expressing Src-GFP (green) and DsRed2-Nuc (magenta) and live imaged. Transgene genotypes of larvae: A58>Src- GFP,DsRed2-Nuc (w1118; +; A58-Gal4, UAS-Src-GFP, UAS-DsRed2-Nuc/+). Projections of a time-lapse series in early L3 larvae. Each frame is merged from 57 planes spaced 0.28 μm apart. Scale bars: 20 μm.

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Kakanj, P., Eming, S.A., Partridge, L. et al. Long-term in vivo imaging of Drosophila larvae. Nat Protoc (2020). https://doi.org/10.1038/s41596-019-0282-z

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