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

Podsumowanie

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

Streszczenie

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

Wprowadzenie

Time-lapse live imaging is a powerful method for studying dynamic cellular processes. The spatial and temporal information provided by time-lapse movies can reveal important details for answering cell biology questions. The Drosophila larva has been a popular in vivo model for investigations using live imaging because its transparent body wall allows for noninvasive imaging of internal structures^1,2. In addition, numerous genetic tools are available in Drosophila to fluorescently label anatomical structures and macromolecules³. However, long-term time-lapse imaging of Drosophila larvae is challenging. Unlike stationary early embryos or pupae, Drosophila larvae move constantly, necessitating immobilization for live imaging. Effective ways of immobilizing live Drosophila larvae include mounting in halocarbon oil with chloroform⁴, anesthetizing using isoflurane or Dichlorvos solution⁵, and compressing between the coverslip and the microscope slide⁶. Although some of these methods have been used for microscopy, none of them is effective for long-term live imaging. Other methods were developed for imaging body wall neurons in crawling larvae using conventional confocal microscopy or light-sheet microscopy^7,8,9. However, these methods are not ideal for monitoring cellular dynamics due to the movement of the larvae.

New methods have been developed to achieve long-term time-lapse imaging of Drosophila larvae. Using a polydimethylsiloxane (PDMS) "larva chip", Drosophila larvae can be effectively immobilized through vacuum-generated suction in a specialized microchamber without anesthetization. However, this method does not offer high temporal resolution for cell biology studies and it has strict limitations on animal size¹⁰. Another method using an anesthetization device achieved live imaging of Drosophila larvae at multiple time points and has been applied to study neuromuscular junctions^{11,12,13,14,15,16}. However, this method also does not allow for continuous imaging for longer than 30 min and requires using desflurane repeatedly, which can inhibit neural activity and affect the biological process studied^17,18. Recently, a new method that combines microfluidic device and cryoanesthesia has been used to immobilize larvae of various sizes for short periods of time (minutes)¹⁹. However, this method requires specialized devices such as a cooling system and longer periods of immobilization require repeated cooling of the larvae.

Here we present a versatile method of immobilizing Drosophila larvae that is compatible with uninterrupted time-lapse imaging for longer than 10 h. This method, which we call "Larva Stabilization by Partial Attachment" (LarvaSPA), involves adhering the larval cuticle to a coverslip for imaging in a custom-built imaging chamber. This protocol describes how to make the imaging chamber and how to mount larvae at a variety of developmental stages. In the LarvaSPA method, the desired body segments are affixed to the coverslip using a UV-reactive glue. A PDMS cuboid additionally applies pressure to the larvae, preventing escape. The air and moisture in the imaging chamber ensure the survival of the partially immobilized larvae during imaging. Advantages of LarvaSPA over other techniques include the following: (1) It is the first method that allows for continuous live imaging of intact Drosophila larvae for hours with high temporal and spatial resolution; (2) The method has fewer limitations on larval size; (3) The imaging chamber and PDMS cuboids can be manufactured at a minimal cost and are reusable.

In addition to describing the larval mounting method, we provide several examples of its application for studying dendrite development and dendrite degeneration of Drosophila dendritic arborization (da) neurons.

Protokół

1. Making the imaging chamber

1. The metal frame can be constructed from an aluminum block in a typical machine shop. The specifications of the frame are illustrated in Figure 1A.
2. To construct the imaging chamber, seal the bottom of the metal frame using a long coverslip (22 mm x 50 mm) and UV glue (Figure 1A). Cure the UV glue using a hand-held UV lamp.

2. Making PDMS cuboids

1. Prepare the mold for PDMS cuboids.
   1. Attach layers of packaging tape to the inner surface of a rectangular (80 mm x 55 mm) Petri dish or round cell culture plate. Use one layer (0.063 mm thick) for second instar larvae, 2 layers (0.126 mm thick) for early third instar larvae, or three layers (0.189 mm thick) for late third instar larvae (Figure 1B).
   2. Cut the tape into strips of specific width with a razor blade: 1.5 mm for the 1-layer tape, and 2 mm for 2-layer or 3-layer tape. The width and thickness of the strip will determine the size of the larvae that the final PDMS cuboid can hold. Leave at least a 5 mm space between the two strips. Remove the tape layers covering the space (Figure 1B).
   3. Remove dust from the inner surface of the plate using sticky tape. The mold is ready for use.
2. Prepare the PDMS mix.
   1. Mix 7 g of PDMS base and 0.7 g of curing agent (10:1 ratio) thoroughly in a small container.
   2. Place the container in a vacuum desiccator for at least 15 min to remove air from the mixture.
   3. Slowly pour about 5.5 g of PDMS mixture onto the mold to reach a 1–2 mm thickness (Figure 1B).
   4. Place the PDMS mixture in the vacuum desiccator again for at least 15 min to remove remaining air bubbles from the mixture. Break the last few bubbles with a pipette tip.
   5. Cure the PDMS on a flat surface in a heat incubator at 65 °C for 2 h.
   6. Use a razor blade to loosen the cured PDMS along the edge of the mold and detach it from the mold. Store the PDMS between two pieces of large sticky tape at room temperature.
   7. For early and late third instar larvae, cut the PDMS into 8 mm x 2 mm x 1 mm cuboids (along the dotted lines in Figure 1B) by positioning the groove created by the tape strip (step 2.1.2) at the center of the long side of the cuboid (Figure 1B,C). For second instar larvae, cut the cuboid to 8 mm x 1 mm x 1 mm.

3. Mounting larvae for long-term time-lapse imaging

1. Prepare the top coverslip for mounting.
   1. Choose six PDMS cuboids with grooves matching the sizes of the larvae. Follow the recommended groove and size of PDMS based on steps 2.1.1, 2.1.2, and 2.2.7.
   2. Remove dust from the surface of the PDMS with sticky tape.
   3. Attach four pieces of double-sided tape (12 mm x 5 mm) on a long coverslip (22 mm x 50 mm) for fixing PDMS cuboids later. The spaces between the two pieces of double-sided tape should be the same as the width of the PDMS groove.
   4. Apply a small drop (~1.2 μL) of UV glue into the groove of each PDMS cuboid and add six small drops of UV glue into the space between the double-sided tape on the coverslip.
2. Prepare the larvae for mounting.
   1. Using a pair of forceps, clean the larvae in water to remove food from the body surface.
   2. Place clean larvae on a small piece of moistened tissue paper in a small (35 mm) Petri dish without a lid. Place the small Petri dish into a large (60 mm) Petri dish containing a piece of dry tissue paper. In a chemical hood, apply 8–12 drops (160–240 μL) of isoflurane onto the dry tissue paper using a plastic transferring pipette and close the lid of the large Petri dish.
   3. Wait 2–3 min while monitoring the larvae. Take out the larvae from the large Petri dish once their mouth hooks stop moving.
3. Mount the animals.
   1. To image structures on the dorsal side of the animal, place the immobilized larvae onto the UV glue between the double-sided tape on the coverslip with the dorsal cuticle facing the coverslip.
   2. Cover each larva with a PDMS block and fit the trunk of the larva into the groove of the PDMS. Leave the head and the tail of the larva outside the PDMS groove. Avoid blocking the spiracles of the larva by the glue.
   3. Press down on the ends of the PDMS block onto the double-sided tape without applying force on the groove. Gently pull on the tail of the larva to flatten the cuticle under the PDMS.
   4. Cure the UV glue for 4 min using a hand-held UV lamp at the high setting (at about 0.07 mW/mm²).
      CAUTION: Protect eyes with safety glasses while using the UV lamp.
   5. Flip the coverslip upside down and repeat step 3.3.4.
   6. Moisten a small piece of lens paper (15 mm x 30 mm) with 20 µL–30 µL of water. Place the moistened lens paper at the bottom of the imaging chamber (Figure 1A,D).
   7. Place the coverslip on the chamber so that the larvae are facing the inside of the chamber. Adhere both ends of the coverslip to the metal surface using UV glue (Figure 1A,D). The dorsal side of the larvae is ready for imaging under confocal microscope (Figure 1E).

4. Imaging

1. Image larvae with an appropriate microscope. All results shown in this protocol (Figure 2, Figure 3, Video S2, Video S3, and Video S4) were acquired using a confocal system with a 40x (1.30 NA) oil objective.

5. Recovery of imaging chamber and PDMS cuboids

1. After imaging, remove the oil on the top coverslip using a lens paper. Detach the top coverslip from the metal frame by cutting into the space between the coverslip and the metal frame with a razor blade. The imaging chamber is ready for reuse.
2. Detach the PDMS cuboids from the top coverslip with forceps. Roll the PDMS cuboids on sticky tape to remove glue residue and dust. The PDMS cuboids are ready for reuse.

Wyniki

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

Dyskusje

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

Materiały

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