Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.


Introduction
The fruit fly, Drosophila melanogaster, has been utilized as a model organism for over 100 years, and has proved instrumental in defining fundamental signaling and developmental pathways that are conserved from invertebrates to human. Live imaging is an important approach to studying cellular mechanisms, and the simple body plan and translucent cuticle of the Drosophila larva makes it an attractive system for live imaging, particularly since there are many genetic tools available for expressing fluorescently tagged proteins in specific cell types.
An important challenge for live imaging techniques is to noninvasively immobilize and position an animal for microscopy. Conventional immobilization approaches include dissection 1,2 or the use of chloroform, both of which kill the animal. The anesthetics ether 4 and isofluorane [5][6][7][8] have also been used. While anesthetics provide many advantages, they also inhibit neural activity and important physiology (including the heartbeat) [9][10][11] , hence can affect the process studied and create stress on the animal. There are also human safety concerns for working with ether and isofluorane.
We have developed a drug-free method to immobilize Drosophila larvae in a single layer PDMS microfluidic device, which we call the 'larva chip' 12 . This protocol will describe how obtain or make the larva chip, and how to utilize it for live imaging in early-staged 3 rd instar larvae.
The chip is comprised of a snug-fitting microchamber, which, upon application of a vacuum via a syringe, immobilizes the animal via gentle mechanical force. The immobilization method brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to a glass coverslip. This allows for high resolution imaging of such structures with high numerical aperture (high magnification) objectives.
13. Switch the valve off such that the PDMS chip is isolated from the syringe and from the environment. As a result, a relatively stable vacuum level is maintained in the microchamber without the need for holding the syringe plunger. 14. Check the larva under the stereoscope to make sure that the entire animal body is placed inside the microchamber, and that the animal is immobile. The trachea should be visible.  Figure 1C). 16. Use a high-magnification objective (oil-immersion, 40-63X is recommended) to locate the animal's structure(s) of interest and perform the imaging. In some cases, a lower magnification may be needed to identify the desired region for imaging before switching to higher magnification. 17. When imaging is completed, release the vacuum by switching the valve to the position that is open to the environment. 18. Detach the PDMS chip from the coverslip. The larva should be immediately motile. 19. Use forceps to remove the larva from the microchamber and gently place the larva on a grape-juice agar plate for recovery.

Inducing a Nerve Crush Injury to Larval Segmental Nerves
1. Follow step 2.3 above to isolate early staged 3 rd instar larvae of the desired genotype. As described in step 2.3 bathe the larvae in water to remove the food. 2. Use a standard fly CO 2 anesthetization station, with CO 2 pad kept under a dissection stereomicroscope, to subdue the larvae. Larvae should become immotile after placement on CO 2 pad for 1-2 min. 3. Now place a single anesthetized larva onto a grape juice agar plate under the stereomicroscope. Turn the animal ventral side up to visualize the ventral nerve cord and segmental nerves through cuticle (Figure 3). Make sure that the larva is completely immotile. 4. Using Dumostar number-5 forceps, pinch the segmental nerves tightly through the cuticle for 5-10 sec. When this is done correctly, the cuticle remains intact and the body wall is not pierced. Note: The injury can be conducted at different positions along the anterior-posterior body axis, as long as the ventral nerve cord, salivary glands, and intestines are not damaged. The most effective injury location is towards the end of the 3 rd abdominal segment, as shown in Figure 3D. Injury at this location damages the most nerves and is the easiest to reproduce without killing the animal. 5. After the injury, turn the animal so that its ventral side down on the grape plate. It should be able to move its head and eat. If the injury was successful, then the posterior half of the larva will be paralyzed. 6. Keep the injured animals on the grape juice agar plate at 25 °C for the desired time according to the experimental objective. For motoneurons, the proximal stump begins to sprout within 8-10 hr of injury 14 , and the distal stump starts to degenerate within 6-8 hr 15 . For class IV da sensory neurons, the proximal stump begins to sprout within 4-6 hr, and the distal stump start to degenerate within 3-4 hr after injury. Note: with appropriate Gal4 drivers and fluorescent reporters, the sprouting and degeneration can be observed in the larva chip (for example, see Figure 6).

Representative Results
The larva chip is comprised of a single layer PDMS block, (a PDMS chip) whose design is outlined in schematic in Figure 1. (See also the supplementary DXF file for designing your own mold). The larva microchamber, vacuum port, and perimeter channels ( Figure 1A) are 140 μm indentations in the PDMS chip. The chip is positioned on top of an early staged 3 rd instar larva, which rests on top of a coverslip with oil ( Figures   1B and 1C). The oil-glass interface between the coverslip and PDMS chip allows for a seal to be created upon application of a mild vacuum. This seal traps the larvae within the chamber, and since the early staged 3 rd instar larva is slightly thicker than the chamber, sealing the chamber creates some physical constriction on the animal, effectively immobilizing and restricting its movement. In this immobilized state, certain ventral body structures, such as the ventral nerve cord and segmental are pushed close to coverslip. This is advantageous for imaging, since in the immobilized state these structures can lie within the working distance of 40X and 63X objectives. After the vacuum is released, the larva can be easily removed from the microchamber, allowing additional experiments to be performed. This purely mechanical immobilization approach can keep 90% of larvae alive after continuous immobilization periods of up to 1 hr 12 .
The vacuum is created by a simple 20 ml syringe, hence the whole unit is easy to transport from a stereomicroscope, where the positioning in the chamber is performed, to a confocal or epifluorescence microscope, where live imaging is performed. The syringe is connected to the vacuum port via polyethylene tubing and 23 G dispensing needles (with lock hubs removed), as described in steps 1.6-1.14. For inverted microscopes, the tubing and syringe are connected via the top of the chip (Figures 1B, 2A, and 2B). For upright microscopes, they are connected via a port on the side of the chip (Figures 1C, 2C, and 2D). The configuration for inverted microscopes is somewhat easier to use. The syringe is pulled to create a gentle vacuum (of approximately 10 psi), which binds the oil-glass-PDMS interface to form a tight seal between the coverslip and the PDMS device, trapping and immobilizing the larva within the chamber.
The placement of the larva into the microchamber (steps 2.7-2.10 in the protocol) is critical for effective immobilization and survival (Figures 2E-H). If the animal is too large for the chamber, (Figure 2G), or if its head or trachea become caught between the edge of the chamber and the coverslip (Figure 1H), then it is unlikely to survive the procedure.
The following are several examples of the use of the larval chip to study various cellular responses in neurons (Figures 4-7, Movie S1 and Movie S2).  axonal transport. This is also sufficient time for desired manipulations in live animals, such as laser based microsurgery, photobleaching, and photoconversion.
To study events longitudinally over a longer time period in a single animal, animals can be placed into the chip and imaged multiple times, separated by periods of rest. Grape juice agar plates are ideal for resting between imaging sessions, as they provide an easy food source and humidity. Multiple imaging sessions do affect larval survival to a degree, since each session carries some risk for damaging the animal (see part 2 in troubleshooting, below). Animals can be routinely imaged >5 times over the course of two days with a greater than 50% survival rate. Since the animals are not anesthetized, they are healthy and motile immediately after release of the vacuum in the chip. There is therefore no need for recovery time between imaging sessions, so the time spacing between sessions is flexible and can be adjusted to the objectives of the experiment.

Troubleshooting:
The most common technical issues with larva chip and recommended solutions are the following: (1) The animal is moving too much. Too much mobility can interfere with the imaging goals. The most common reasons for this in the larva chip are a) the animal is too small for the chip, or b) the vacuum pressure applied during the immobilization step is compromised. The larva chip described in this protocol is designed for early staged 3 rd instar larvae. The optimal size for the animal is 3.5-4 mm in length (along the anteroposterior axis). To ensure that the vacuum pressure is sufficient, pull the syringe 2-2.5 ml, or until resistance is felt in the handle. One indication that the vacuum is working is that small bubbles in the perimeter channel may be seen moving slowly towards the vacuum source. Another indication is that the coverslip should always travel with the chip when the chip is lifted from the top (and this is the recommended method for transporting the chamber once the larvae is positioned and the vacuum is on). The vacuum may be compromised if there are cracks in the tubing, or if there is oil in the tubing. This can be easily addressed by replacing the 23 G dispensing needle tip and polyethylene-50 tubing (from steps 1.6-1.14).
(2) The animal dies after imaging in the chip. The procedure is intended to cause minimal stress upon the animal, and animals of wild type genotype have a >90% survival rate, even after an hour of immobilization on the chip 12 . Since some genotypes may be less resilient to the stress of the chip, first check that wild type animals (for example, Canton S) survive the immobilization technique. a) The most common cause for lethality is incorrect positioning of the larva (see Figures 2G-H). If parts of the cuticle, head or trachea are not entirely within the chamber, then they can become damaged during the immobilization, and a larva that is too large for the chip (>4 mm) is less likely to survive. b) A less common cause for lethality is the use of too much pressure or vacuum when loading the chip. When properly positioned in the chip, the pressure generated by the vacuum is well tolerated. However excessive pressure, either from the vacuum or in the initial stage of positioning the animal can be an issue. It is best to learn the degree of pressure needed empirically by trials with wild type larvae of the correct size. c) If too much Halocarbon oil covers the animal's trachea the animal may potentially have issues with long-term survival. The oil plays several important roles in the chip: it is important for creation of the vacuum, the optics during imaging, and it counteracts desiccation in the chip. However excessive oil should be avoided. (This can also lead to oil in the tubing and syringe, compromising the vacuum). The suggested protocol coats just the ventral side of the larva with oil, then removes excess oil by placement of the larva on a clean coverslip before transferring to the final coverslip for imaging. d) phototoxicity can be experienced from the imaging session. As with any live imaging application, it is ideal to use short exposure times with low intensity laser light, which is best achieved using a highly sensitive camera or detector. Try to minimize illumination with UV light, including broad-spectrum light created by Hg light sources.

Other issues and future directions:
Since this method does not utilize anesthetics, the animal's heart continues to beat. This creates some unavoidable mobility, which affects imaging in some locations more than others. The examples here demonstrate that the ventral nerve cord, segmental nerves, and body wall can be readily imaged without interference from the heartbeat. In cases where the heartbeat affects imaging, the regular movements can sometimes be corrected for within analysis software (for example, the Image Stabilizer plugin for ImageJ). This works well when individual objects are moving on a fast time scale (for example ~1 μm/sec for fast axonal transport) or on a very slow time scale (minutes to hours). However, when the object(s) of interest move with a range of speeds and directions, it can be harder to correct for the heartbeat induced movements.
Another issue is slight variability in optics from animal to animal, or between multiple imaging sessions of the same animal in the chip. The deeper the object of interest is within the animal, the greater this variation will be. Segmental nerves and the ventral nerve cord are normally too deep within then animal to be imaged on a regular microscope. However the mild pressure experienced in the larva chip pushes these structures very close to the cuticle and coverslip. The exact distance of these structures from the coverslip will have small variations from trial to trial. The variation for objects close the cuticle, such as the cell bodies of sensory neurons, is less. It is therefore important, particularly for making measurements of intensity, to utilize a large number of animals and independent trials to account for the variability in optics.
While the examples shown here have focused upon processes within neurons, the approach should be amenable to imaging any structure in the animal that can be brought within the focusing depth of the microscope objective. This includes the cuticle, body wall muscles, and their NMJs. Trachea on the ventral side of the animal and potentially parts of the digestive tract might also be imaged. The animal may also be positioned with its dorsal side towards the coverslip for short term imaging of structures near the dorsal surface. The ability to image structures deep within the animal is limited by the working distance of the microscope objective used. Structures such as imaginal discs are inaccessible to high magnification (e.g. 40X) objectives.
The larva chips described in this protocol are designed for larvae in the early 3 rd instar stage (ranging in size from 3.5-4 mm). However many interesting questions require imaging at different larval stages. Smaller chips to accommodate 2 nd instar larvae, or larger chips to accommodate late 3 rd instars can be easily designed using the same principle. (Supplementary Figure 1 contains a readily modifiable DXF file for making