The Photoconversion Technique for Exploring Inflammatory Cell Dynamics in Insect Pupae

This protocol consists of four main sequential steps: (1) Preparation of Drosophila stocks and staging of Drosophila pupae, (2) Pupal dissection and mounting, (3) Pupal wounding, (4) in vivo time-lapse confocal imaging.

1. Preparation of Drosophila Stocks and Staging of Pupae

1. Obtain appropriate Drosophila stocks (see Introduction and Table of Materials).
2. Collect young healthy adult flies of the appropriate genotype.
   1. Select adult flies by using carbon dioxide gas pads to briefly anesthetize the flies and a fine paintbrush to transfer flies of the appropriate genotype or gender to a collection vial.
   2. Add 20 virgin females and 20 males to each vial containing standard fly food media (a cornmeal-molasses-agar mixture, see Table of Materials) supplemented with yeast.
3. For optimal pupal generation, tip the adult flies each day onto new food in fresh vials, keeping all vials at 25 °C.
   NOTE: If the Gal4-upstream activating sequence (Gal4-UAS) system is being used to drive lineage-specific gene expression, all steps should be performed at 25 °C or above, as the Gal4-UAS system is temperature sensitive.
4. 18 h before the scheduled imaging session, select at least 10 newly formed white pre-pupae (Figure 1A) from the vials (i.e. 0 h after puparium formation, APF) using forceps or a fine paintbrush to dislodge pupae from the interior vial surface and transfer pupae carefully to the side of a clean empty plastic vial.
   NOTE: Wandering 3^rd instar larvae crawl upwards out of the food medium to undergo pupation; newly formed white prepupae are easily identified as they possess everted anterior spiracles and are stationary (unlike 3^rd instar larvae). The cuticle transforms into the 'puparium' (the pupal case) which is initially soft and white. Care must be taken to avoid damaging the pupae, as this can lead not only to an unwanted injury-induced inflammatory response but also to a significant developmental delay.
5. Age the selected pupae to the appropriate developmental stage (18 h APF will give optimal results) in the vial at 25 °C.
   NOTE: As the pupae develop, the pupal case will become progressively darker and more brittle.
6. Prepare other reagents for the next step ahead of time. To make the heptane glue, combine a 20 cm length of rolled-up double-sided tape with 20 mL heptane in a 50 mL centrifuge tube, seal with the paraffin film, and rock at room temperature overnight on the bench.

2. Preparation and Dissection of Drosophila Pupae

1. Transfer the staged Drosophila pupae to a piece of double-sided sticky tape mounted on a glass slide. Position the pupae so that the ventral side is firmly stuck to the tape and the dorsal side is facing upwards (Figure 1B).
   NOTE: The anterior of the pupa will be identifiable from the two spiracles protruding from the anterior end of the pupal case.
2. Carefully remove pupae from their protective puparium casing under a brightfield dissection microscope using forceps and micro-scissors (Figure 1B-D).
   1. Initially, make an incision in the anterior-most region of the puparium using the forceps (Figure 1B). Ensure that the pupal case in this area is hollow and devoid of pupal tissue because the pupae will have shrunk within the case during early pupal development.
   2. After this initial incision, carefully tear or cut the pupal case open in an anterior-to-posterior direction using forceps or microscissors (Figure 1C) until the pupa is completely free from the brown opaque brittle casing (Figure 1D).
      NOTE: Pupae at this stage are very fragile and care must be taken to avoid puncturing the pupal surface; puncturing is obvious as hemolymph quickly leaks out from the puncture site. Pupae with punctures, however small, should be discarded.
3. Mount the pupae in a glass-bottomed dish using heptane glue.
   1. Use a 20 mL pipette tip to place a 10 mL drop of pre-prepared heptane glue (see section 1.6 above) in a line on the glass-bottomed dish.
   2. Allow the glue to dry for 5 s before transferring the dissected pupae carefully onto the heptane glue using forceps (Figure 1E).
   3. For ease of wounding and imaging, line the pupae up in a row; approximately 5 pupae are suggested but more will be manageable with experience.
      NOTE: The use of heptane glue is recommended when using upright imaging systems but is not necessary when using an inverted system. If the glue creates optical aberrations during imaging, pupae can be placed directly on the coverglass instead – the natural adhesion between the pupal tissue and glass will be sufficient in most cases for stable imaging, as long as care is taken when moving the imaging dish between microscopes.
4. For best results, mount the pupae so that the wing is flat on the coverglass with the majority of the wing surface in direct contact with the coverglass (Figure 1F). Roll the pupae using forceps to change their position to ensure the wing is mounted correctly.
5. To prevent sample dehydration during the imaging period, add a piece of absorbent filter paper soaked in distilled water to the side of the glass-bottomed dish at the end of mounting, being careful not to disturb the pupae (Figure 1E). Cover the dish with a lid.  
   NOTE: Pupae are now ready for wounding and imaging.

3. Laser-induced Wounding of Drosophila Pupal Wings

1. Transfer the glass-bottomed dish containing the mounted pupae to a wide-field microscope equipped with a tunable laser ablation system.
   1. Use a pulsed-UV air-cooled nitrogen-pumped ablation laser tuned to 435 nm – see Table of Materials for details; the precise wavelength of the light used for illumination is selected by the user via an appropriate dye cell.
2. Using brightfield optics, adjust the microscope stage controls to locate the pupal wing of the first pupa to be wounded (Figure 1F).
3. For optimal results, use an oil immersion 40X or 63X objective lens for both the imaging and laser ablation; ensure that the immersion liquid used (oil or glycerol) is consistent between ablation and imaging systems. Adjust the microscope to focus on the plane of the pupal wing epithelium nearest the glass coverslip (i.e. focus on the region of the epithelium to be wounded) using the fine focus control knob.
4. Using the microscope stage adjustments, position the pupal wing so that the area to be wounded is directly aligned with the known target area of the ablation laser.
5. Use an energy density attenuator slide fitted to the microscope to manually adjust the power level of the ablation light.
   NOTE: The attenuator slider has click stops and rulings to identify the relative level of attenuation and permit the use of reproducible settings.
6. Using an external manual trigger control, activate the ablation laser using a single brief click of the trigger to make a wound. Check for the appearance of the transient air bubble at the ablation site since wounding will normally be accompanied by it. Check if the laser-induced wounding has been successful using the appropriate fluorescent filters to visualize the pupal epithelium.   
   NOTE: Take care to avoid accidental exposure to the laser beams as the beam reflections can cause severe eye or skin damage.
7. If wounding is unsuccessful, vary the focal plane (moving the microscope focus slightly above or below the current focal level) and repeat the single click of the ablation trigger. Alternatively, gradually increase the laser power using the attenuator slide until the desired wound size is achieved.
8. To vary the ablation laser pulse repetition rate, use the Repetition rate knob on the rear control panel (which changes the rate from less than 1 pulse/s up to 60 Hz). For optimal wounding, set the pulse repetition rate to 40 Hz.
9. To generate different-sized wounds, use the energy density attenuator slide to manually adjust the power level of the ablation light.
10. Refrain from wounding all of the mounted pupae and use these non-ablated pupae as unwounded controls.
11. For consistent results, regularly realign the ablation system (using the relevant operating manual). Also, clean and refill the dye resonator cell that controls the laser output wavelength.

4. In Vivo Time-lapse Confocal Imaging

1. Quickly transfer the glass-bottomed dish to an appropriate microscope for time-lapse imaging.       
   NOTE: For optimal results, use a high-specification confocal or spinning disc microscope equipped with sensitive detectors that can detect both green fluorescent protein (GFP) and mCherry fluorophores.
2. To image the entire pupal wing, use a low magnification (e.g. 20X) objective lens (Figure 2A). To image wound repair and the accompanying inflammatory response with high spatial resolution, use the oil immersion 40X (NA 1.3) or 63X (NA 1.4) objective lenses (see representative images in Figure 2B-D).
3. Open the appropriate image capture software associated with the microscope.
4. Using the image capture software, turn on the appropriate lasers e.g. the 488 nm and 561 nm lasers to visualize GFP and mCherry fluorophores, respectively (by clicking in the relevant boxes) and adjust the laser power and gain/offset settings to give sufficient fluorescent signal whilst avoiding pixel saturation; use the lowest possible laser power (in the range 5 – 20%) to minimize photobleaching and phototoxicity.
5. To capture both the repairing epithelium and inflammatory cell recruitment, set the microscope to record a z-stack using the fine focus adjustment knobs on the control panel; for optimal results, set the software (using manual button-clicks) to record z-slices through the pupal wing (minimum every 3 mm), from the top of the wounded epithelium through to the extracellular space beneath (containing migrating hemocytes) to achieve a large z-stack (in the range of 50 - 100 mm).
6. For time-lapse imaging, record z-stacks at regular time intervals (minimum every 30 s) for at least 1 h post-wounding. 
   NOTE: The exact time interval between the z-stacks chosen represents a trade-off between capturing the rapidly changing cell dynamics and avoiding photo-bleaching of the samples.
7. To simultaneously image multiple pupae (including non-ablated unwounded controls), use a motorized stage (attached to the microscope) and the multi-position acquisition feature available within the imaging software. Manually set the position of each pupa within the software using the stage position control knobs and then manually set the appropriate z-stack limits (top and bottom) for each pupa.
8. Visualize the time-lapse images either during image capture or later with specialist image analysis software (such as ImageJ) using z-stack projections or 3-D rendering. For example, to follow the movements of individual hemocytes (as in Figure 2C' and D'), track hemocyte nuclei using the open-access ImageJ plug-ins "TrackMate" or "Manual Tracking" (methods published in).
9. Use photoconvertible probes (such as Kaede) to selectively photoconvert and label a subset of epithelial or immune cells during imaging.
   1. Open appropriate modules within the imaging software to perform the photoconversion (such as the Fluorescence recovery after photobleaching (FRAP), and fluorescence recovery after photo-bleaching module) and activate the 405nm laser (by clicking in the relevant software box).
   2. Select cells to be photoconverted within the FRAP software using the square, circular, or freehand selection tool. Within the FRAP software, set the time course for photoconversion (Bleaching) to a single iteration/frame and set the 405 nm laser at 20% laser power. Manually click Start the experiment to perform photoconversion.
   3. Exit the FRAP module (click Close) and return to the original imaging screen within the software; use the 488 nm and 561 nm lasers to image the behavior of photoconverted and non-photoconverted cells by setting up a z-stack and time-lapse recording as above.
      NOTE: Photoconverted probes remain stable for many hours after the initial photoconversion, enabling the behavior of the photoconverted cells to be followed over time (for at least 5 h). For example, inflammatory cells in the wound can be selectively photoconverted (Figure 2F) and their behavior followed as they resolve from the injury site (Figure 2G and H).