This work was supported by the National Science Foundation, (grant number IOS-0842701 to C.A.C), and the National Institute of Health (R00MH080599 to B.Y., R21 NS062313 to N.C, and NS069844 to C.A.C). We would like to acknowledge James Schutt, Emily Han, and Leni Truong for technical support, and the Bloomington Stock center for fly lines. All the chips were fabricated at the Lurie Nanofabrication Facility at the University of Michigan.

1. Making the PDMS Chip
To make a PDMS chip from the SU-8 mold, follow steps 1.1-1.7. If a chip is on-hand, but needs to assembled for use, skip to step 1.8.
<ol>
	<li>Mix 45 g of PDMS base and 4.5 g of curing agent (10:1 ratio) from a PDMS kit in a small disposable plastic container and mix them thoroughly using a plastic stir stick.</li>
	<li>Place the container in a vacuum container (e.g. a desiccator) for 10 min to remove any bubbles.</li>
	<li>Place the SU-8 mold at the bottom of a 150 mm in diameter plastic dish and slowly pour the PDMS mixture on the mold. Take care not to generate bubbles while pouring the PDMS.</li>
	<li>Cure the PDMS in an oven (or incubator) at 650 &#176;C for 4 hr.</li>
	<li>Remove the cured PDMS/SU-8 mold from the oven and let it cool for a few minutes.</li>
	<li>Using a razor blade, cut the cured PDMS along the edge of the SU-8 mold and detach it from SU-8 mold.</li>
	<li>Divide the PDMS slab into individual PDMS chips using a razor blade.</li>
	<li>Using a 21 G&#160;dispensing needle, poke a hole in the vacuum port (depicted in Figure 1A) of the PDMS chip.</li>
	<li>Take a 23 G dispensing needle and twist the needle tip from its base a few times to break the needle tip off from the lock hub.</li>
	<li>Insert the 23 G needle tip into a small piece of polyethylene tubing so that the tubing covers at least a millimeter of the needle. Then use a razor blade to cut the excess tubing away from the needle. This creates a plastic ring around one end of the needle, which will create a seal when inserted into the vacuum intake port.</li>
	<li>For use with an inverted microscope (Figures 1B and 2A-B): insert the 23 G needle tip into the hole of the vacuum port. For use with an upright microscope (Figures 1C and 2C-D): poke a second hole on the side of the PDMS chip with a 21 G dispensing needle; this hole will provide access to the first hole from the side. Then insert the 23 G needle tip with tubing ring into the side hole. Place a piece of double-sided tape over the top of the PDMS chip to seal the top hole (Figure 1C).</li>
	<li>Take a piece of polyethylene tubing that is approximately 20 cm in length. Connect one side of the tubing to the needle tip that is inserted into the vacuum port.</li>
	<li>Connect the other side of the tubing to one of the ports of a 3-way valve (see '3-way stopcock' in the list of materials)</li>
	<li>Attach a 20 ml syringe into one of two remaining ports. The last port is open to the environment.</li>
</ol>
2. Using the Larva Chip for Live Imaging
<ol>
	<li>Clean the PDMS chip with transparent adhesive tape. Attach a piece of tape to the bottom of chip. Make sure the tape is touching the entire PDMS surface, and then peel off the tape.</li>
	<li>Repeat the above step 2-3x to make sure there are no particles or oil (retained from previous experiments) on the surface of the PDMS chip. Since the PDMS chip is reusable, it is very important to remove oil residue as it can affect the adhesion of PDMS to glass and result in inadequate sealing.</li>
	<li>Transfer early (i.e. foraging) 3rd instar larvae to a Petri&#160;dish containing water. (The foraging 3rd instar stage larvae are in the food, rather than the side of the culture vial). Bathe the larvae in water to remove culture medium.</li>
	<li>Take a clean glass coverslip and place a small drop of Halocarbon 700 oil at its center.</li>
	<li>Using forceps, gently pick up a clean, early-staged 3rd instar larva from the water (the larva should be ~3.5-4 mm in length). Place the animal briefly on a lightweight wipe or paper towel to remove excess water, and then place it on the oil drop. The drop should be small enough such that the trachea of the larvae are not coated. Let the larva stay on the oil drop for 10 sec.</li>
	<li>Remove the larva from the oil drop and then place it on a clean glass coverslip.</li>
	<li>Transfer the larva to another clean glass coverslip. This step removes excess oil.</li>
	<li>Pay attention to larva orientation. For imaging the neural cord and segmental nerves, the ventral side of larva should sit on the coverslip. Its dorsal side, characterized by two longitudinal tracheal tubes, should face upwards. Note: this is the orientation the larva prefers naturally.</li>
	<li>Gently place the PDMS chip on top of the larva. The larva should be aligned to center middle of the microchamber, with its tail oriented towards the vacuum port. Be careful that the larva does not touch the edges of the chamber. This is especially important for the anterior and posterior tracheal terminals. Note: this step is best done under a stereomicroscope.</li>
	<li>Push the PDMS chip against the glass coverslip to achieve a good seal. Ensure that the larva is entirely enclosed by the microchamber when the PDMS chip is touching the glass coverslip.</li>
	<li>Switch the 3-way valve on such that the syringe can draw air from the PDMS microchamber (through the tubing) to create a vacuum.</li>
	<li>With one hand, hold the PDMS chip/glass coverslip firmly. Use the other hand to pull the syringe plunger. Withdraw 2-2.5 ml of air, until resistance is felt in the syringe handle, to create vacuum. The vacuum produces a tight seal between the PDMS chip, oil, and coverslip interfaces and restricts the mobility of the larva.</li>
	<li>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.</li>
	<li>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. The rest of the PDMS chip should be in contact with the coverslip. Note: See Figures 2E and 2F for examples of animals correctly immobilized in the chip. Some incorrect orientations are shown in Figures 2G and 2H.</li>
	<li>Place the larva chip (PDMS chip+ glass coverslip) on the microscope. The larva chip, the tubing and the syringe should be handled carefully to avoid detachment of the PDMS chip from the coverslip. For an upright microscope, fix the 'top' side of the chip to the microscope stage with double-sided tape (Figure 1C).</li>
	<li>Use a high-magnification objective (oil-immersion, 40-63X&#160;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.</li>
	<li>When imaging is completed, release the vacuum by switching the valve to the position that is open to the environment.</li>
	<li>Detach the PDMS chip from the coverslip. The larva should be immediately motile.</li>
	<li>Use forceps to remove the larva from the microchamber and gently place the larva on a grape-juice agar plate for recovery.</li>
</ol>
3. Inducing a Nerve Crush Injury to Larval Segmental Nerves
<ol>
	<li>Follow step 2.3 above to isolate early staged 3rd instar larvae of the desired genotype. As described in step 2.3&#160;bathe the larvae in water to remove the food.</li>
	<li>Use a standard fly CO2 anesthetization station, with CO2 pad kept under a dissection stereomicroscope, to subdue the larvae.&#160;Larvae should become immotile after placement on CO2 pad for 1-2 min.</li>
	<li>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.</li>
	<li>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 3rd 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.</li>
	<li>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.</li>
	<li>Keep the injured animals on the grape juice agar plate at 25&#160;&#176;C for the desired time according to the experimental objective. For motoneurons, the proximal stump begins to sprout within 8-10 hr&#160;of injury14, and the distal stump starts to degenerate within 6-8 hr15. 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&#160;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).</li>
</ol>

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The authors declare no competing financial interests.

Making or obtaining the larva chip:
The larva chip consists of a PDMS block (termed the 'PDMS chip') attached to a glass coverslip. The protocol in step 1 describes the procedure for making and using larva chips, assuming a SU-8 mold is available. The SU-8 mold is microfabricated by photolithographically patterning a 140 &#956;m thick SU-8 photoresist layer on a silicon wafer (for details see Ghannad-Rezaie&#160;et al.12). As the microfabrication of the SU-8 mold requires access to specialized equipment, we recommend ordering it from a microfabrication facility (e.g. the LNF facility at the University of Michigan14), or from a foundry by sending them the chip design that is provided as a supplementary file.&#160;If one wishes to change the design of the PDMS chip (e.g. for use with larvae of different sizes), a CAD software that handles DXF files (e.g. Autocad) can be used. An SU-8 mold can also be made in-house following instructions in Mondal&#160;et al.27&#160;Many readers may find it convenient to simply obtain a sample PDMS chip to try out the technique before fabricating their own chips. This will be made freely available upon request.
Use of the microfluidic &#39;larva chip&#39;&#160;for live imaging:
The immobilization method in the larva chip avoids the use of anesthetics, and instead involves pressure, via the application of a vacuum, to restrict the animal's movement. While animals can survive immobilization in the chip for multiple hours12, a shorter immobilization period (5-15 min) is recommended. This is enough time for imaging many cellular events of interest, including changes in intracellular calcium, or fast 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 &#62;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 3rd 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 &#62;90% survival rate, even after an hour of immobilization on the chip12. 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 (&#62;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.&#160;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.&#160;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 &#956;m/sec&#160;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.&#160;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 3rd instar stage (ranging in size from 3.5-4 mm). However many interesting questions require imaging at different larval stages. Smaller chips to accommodate 2nd instar larvae, or larger chips to accommodate late 3rd instars can be easily designed using the same principle. (Supplementary Figure 1 contains a readily modifiable DXF file for making silicon molds with altered chamber sizes). The simple principle of the reversible seal could even be applied to other organisms such as C. elegans or zebrafish, with the main variant being the chamber size. A useful future direction is to design a chip that can immobilize many animals at once, to use for screening purposes.&#160;However, for this, the design would need to be significantly different from the current device, where the issues of positioning the animal in the chip needs to be dealt with for each animal independently.
The nerve crush assay for studying injury responses in larval peripheral nerves:
The nerve crush assay described here for larval segmental nerves is a simple method for introducing an injury to peripheral axons in Drosophila. Advantages of this method include: a) it is simple to conduct with standard tools found in a Drosophila lab (a stereomicroscope CO2 source and forceps); b) it can be conducted quickly for many animals, making biochemical analysis of nerve cords after injury feasible14; c) the molecular and cellular responses to this injury are highly reproducible14,15,28 and can be used to discover processes that are also important in vertebrate neurons29,30.
Alternate methods for injuring neurons is to focus a high-power laser, for example a pulsed-UV or femtosecond laser, to sever an axon via laser microsurgery17,31-33. The larva chip is an ideal method for positioning the animal for such microsurgery. However, because of minor differences in optics between trials, discussed above, the laser-based method can be more difficult to reproduce in larvae, particularly in larval segmental nerves. Also, laser based axonal injury requires more time to position each animal, hence is more difficult to conduct on a large scale (with a large number of animals).
Troubleshooting:
The most commonly encountered technical issue from the nerve crush is death from damage to internal organs. When conducting the crush, it is important not to pinch the ventral nerve cord, salivary glands, or intestines. It is also important not to puncture the cuticle. These issues are best avoided by bringing the forceps at a 45&#176; angle to the cuticle surface (see Figure 3).
The quality of the forceps has a big impact upon the effectiveness of the crush and survival afterwards. We recommend Dumostar number 5 forceps. To retain their sharpness, the forceps must be handled with care, not used for other purposes, and replaced once they become blunt or bent.
The size of the animal can also influence the effectiveness of the crush. Small animals (less than 3 mm in length) are much less likely to survive the injury. With large animals, (wandering 3rd instars), it is more difficult to locate the nerves and avoid damage to the larger salivary glands and intestines, and there is less time to study injury responses before pupation. The nerve crush is most effectively conducted in early 3rd instar larvae (which are ~3-4.5 mm in length along the anteroposterior axis).
The food source that the animal is raised upon may affect the strength of the cuticle and survival after the crush. It is recommended to raise animals in food made from a standard yeast-glucose recipe.
The best method for learning how to do the crush effectively is to practice on many animals, first with the primary goal of achieving survival (and not pupation) 24 hr after the crush. Beginners normally have a low survival rate (e.g. 10%), but once the technique is learned, survival rates can reach ~90%.
Other issues and future directions:
The crush assay provides a powerful method to study the sprouting of axon proximal to the injury site and degeneration of axons and synapses distal to the injury site. While the rates of degeneration vary between different neuron types, they are highly reproducible within a given neuron type, providing testament to the reproducibility of the injury assay.
In contrast, the 'regenerative' sprouting response observed in proximal axons is more challenging to study. All axons in the segmental nerve initiate extensive sprouting close to the injury site (for example, see Figure 6 and Figure 3). However the extent of sprouting can vary from neuron to neuron, and is difficult to quantify.&#160;A similar degree and variability in sprouting can be observed after more focal lesions of single motoneurons in segmental nerves introduced by using a UV pulsed dye laser. We interpret that the nondiscriminate directionality of the sprouting is due to the absence of guidance cues in the segmental nerves. In contrast, sensory neuron axons injured by laser close to their cell bodies undergo new axonal growth in the same direction as the lost axon34. Axons in this region of the animal are likely exposed to more specific positional information for guidance of the regenerating axons. The environment within segmental nerves is unlikely to have much resemblance to the environment that the axons originally navigated during their guidance in the embryo, hence is not expected to have information to guide regenerating axons.
Another limitation for studying regeneration using the segmental nerve crush assay is that injured sensory and motoneuron axons still have a significant distance to cover (0.25-1 mm) to reach their target, and a limited time frame (&#60;3 days) before the animal undergoes pupation. A recent study has identified a genetic manipulation of the prothoraciotropic hormone receptor which triples the duration of the 3rd instar larval stage35. This manipulation will extend the time frame for studying the recovery and degeneration of neurons after injury significantly, to 9 instead of 3 days. This may be long enough to observe new events, such as reconnection of an injured axon with its postsynaptic target, especially if the injury is induced close to the synaptic ending.

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 dissection1,2 or the use of chloroform, both of which kill the animal. The anesthetics ether4 and isofluorane5-8 have also been used. While anesthetics provide many advantages, they also inhibit neural activity and important physiology (including the heartbeat)9-11, 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.&#160;This protocol will describe how obtain or make the larva chip, and how to utilize it for live imaging in early-staged 3rd 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.
Advantages of the larva chip over other conventional techniques include the following: (i) Use of the larva chip replaces the use of chemicals, allowing for in&#160;vivo imaging of unanesthetized animals. (ii) Larvae recover immediately after release from the chip (in contrast to a 2 hr recovery period for isofluorane8,13). This allows for imaging over broad time scales, ranging from milliseconds to minutes, to hours and days. (iii) The use of PDMS, which is a gas permeable material, enables for continuous diffusion of oxygen/air from the environment into the larva body. (iv) The chip is easy and safe to use, and (v) it is reusable, and can be manufactured at a minimal cost.
In addition to instructions for using the larva chip, this protocol will provide several examples of its use to study neuronal events in 3rd instar larvae. These include live imaging of axonal transport, 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 neuronal responses to axonal injury. For this an additional procedure is described (in part 3) for injuring axons within peripheral nerves by a segmental nerve crush. This simple assay can be performed both rapidly and reproducibly under a standard dissection stereomicroscope, which allows for many animals to be processed at the same time. Cellular responses to the injury can be studied by live imaging in the larva chip.

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	<tbody>
		<tr height="38">
			<td height="38" style="height:39px;width:259px;">0.5 mm Polyethylene tubing</td>
			<td style="width:171px;">Fisher scientific</td>
			<td style="width:291px;">14-170-11B</td>
			<td style="width:449px;">Polyethylene tubing, I.D. = 0.023&#8217;&#8217;, O.D. = 0.038&#8217;&#8217;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">1 mm Polyurethane tubing</td>
			<td style="width:171px;">Fisher scientific</td>
			<td style="width:291px;">BB521-63</td>
			<td style="width:449px;">Polyurethane tubing, I.D. = 0.063&#8217;&#8217;,&#160; O.D. = 0.125&#8217;&#8217;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">barb to barb connector</td>
			<td style="width:171px;">Bio Rad</td>
			<td style="width:291px;">732-8300</td>
			<td style="width:449px;">0.8 mm barb to barb connector</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">3-way stopcock valve</td>
			<td style="width:171px;">Bio Rad</td>
			<td style="width:291px;">732-8104</td>
			<td style="width:449px;">Screw on valve for the syringe</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">Syringe (20 ml)</td>
			<td style="width:171px;">Fisher scientific</td>
			<td style="width:291px;">14-817-33</td>
			<td style="width:449px;">Screw on 20 ml syringe for&#160; generating vacuum</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:259px;">Dispensing needles, 23 G (.4mm I.D., .6mm O.D.)</td>
			<td style="width:171px;">McMaster-Carr</td>
			<td style="width:291px;">75165A684</td>
			<td style="width:449px;">Needle for outlet connection</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:259px;">Dispensing needles, 21 G, (.6mm I.D.,&#160; .8mm O.D.)</td>
			<td style="width:171px;">McMaster-Carr</td>
			<td style="width:291px;"><a href="http://www.mcmaster.com/#75165A679" title="Close">75165A679</a></td>
			<td style="width:449px;">Needle for outlet connection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">Halocarbon oil</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:291px;">H8898</td>
			<td style="width:449px;">Halocarbon oil 700</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">Dumostar Number 5 Forceps</td>
			<td style="width:171px;">Roboz</td>
			<td style="width:291px;">RS-498</td>
			<td style="width:449px;">For nerve crush</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:259px;">PDMS Kit (Base and curing agent)</td>
			<td style="width:171px;">Ellsworth</td>
			<td style="width:291px;">184 SIL ELAST KIT 0.5KG</td>
			<td style="width:449px;">Dow Corning Sylgard 184 Silicone Encapsulant 0.5 kg Kit Clear</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:259px;">Glass Coverslips</td>
			<td style="width:171px;">Fisher scientific</td>
			<td style="width:291px;">12-544-C</td>
			<td style="width:449px;">24 x 40 mm (thickness according to recommendation for your microscope objective)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">Disposable Plastic Cup (9 oz.)</td>
			<td style="width:171px;"></td>
			<td style="width:291px;"></td>
			<td style="width:449px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">Plastic coffee stirrer stick</td>
			<td style="width:171px;"></td>
			<td style="width:291px;"></td>
			<td style="width:449px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:259px;">Razor Blade</td>
			<td style="width:171px;"></td>
			<td style="width:291px;"></td>
			<td style="width:449px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:259px;">Grape juice agar plates</td>
			<td style="width:171px;"></td>
			<td style="width:291px;"></td>
			<td style="width:449px;"><a href="http://cshprotocols.cshlp.org/content/2007/4/pdb.rec10925">See http://cshprotocols.cshlp.org/content/2007/4/pdb.rec10925 for recipe</a></td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae at JoVE.com

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

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

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 ...

using-microfluidics-chips-for-live-imaging-study-injury-responses

Research

JoVE Journal

Bioengineering

15.4K Views.  University of Michigan. Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the 'larva chip' for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

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

Bioluminescence Imaging for Assessment of Immune Responses Following Implantation of Engineered Heart Tissue (EHT)

Various techniques of cardiac tissue engineering have been pursued in the past decades including scaffolding strategies using either native or bioartificial scaffold materials, entrapment of cardiac myocytes in hydrogels such as fibrin or collagen and stacking of myocyte monolayers 1. These concepts aim at restoration of compromised cardiac function (e.g. after myocardial infarction) or as experimental models (e.g. predictive toxicology and substance screening or disease modelling). Precise monitoring of cell survival after implantation of engineered heart tissue (EHT) has now become possible using in-vivo bioluminescence imaging (BLI) techniques 2. Here we describe the generation of fibrin-based EHT from a transgenic rat strain with ubiquitous expression of firefly luciferase (ROSA/luciferase-LEW Tg; 3). Implantation is performed into the greater omentum of different rat strains to assess immune responses of the recipient organism following EHT implantation. Comparison of results generated by BLI and the Enzyme Linked Immuno Spot Technique (ELISPOT) confirm the usability of BLI for the assessment of immune responses.

Bioluminescence Imaging for Assessment of Immune Responses Following Implantation of Engineered Heart Tissue EHT

This video demonstrates the use of in vivo bioluminescence imaging to study immune responses after implantation of Engineered Heart Tissue (EHT) in rats.

Various techniques of cardiac tissue engineering have been pursued in the past decades including scaffolding strategies using either native or ...

Impulsive Pressurization of Neuronal Cells for Traumatic Brain Injury Study

A novel impulsive cell pressurization experiment has been developed using a Kolsky bar device to investigate blast-induced traumatic brain injury (TBI). We demonstrate in this video article how blast TBI-relevant impulsive pressurization is applied to the neuronal cells in vitro. This is achieved by using well-controlled pressure pulse created by a specialized Kolsky bar device, with complete pressure history within the cell pressurization chamber recorded. Pressurized neuronal cells are inspected immediately after pressurization, or further incubated to examine the long-term effects of impulsive pressurization on neurite/axonal outgrowth, neuronal gene expression, apoptosis, etc. We observed that impulsive pressurization at about 2 MPa induces distinct neurite loss relative to unpressurized cells. Our technique provides a novel method to investigate the molecular/cellular mechanisms of blast TBI, via impulsive pressurization of brain cells at well-controlled pressure magnitude and duration.

A novel impulsive cell pressurization experiment has been developed using a Kolsky bar device to investigate the molecular/cellular mechanisms of blast-induced traumatic brain injury.

A novel impulsive cell pressurization experiment has been developed using a Kolsky bar device to investigate blast-induced traumatic brain injury (TBI). ...

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

Endothelialized Microfluidics for Studying Microvascular Interactions in Hematologic Diseases

Advances in microfabrication techniques have enabled the production of inexpensive and reproducible microfluidic systems for conducting biological and biochemical experiments at the micro- and nanoscales 1,2. In addition, microfluidics have also been specifically used to quantitatively analyze hematologic and microvascular processes, because of their ability to easily control the dynamic fluidic environment and biological conditions3-6. As such, researchers have more recently used microfluidic systems to study blood cell deformability, blood cell aggregation, microvascular blood flow, and blood cell-endothelial cell interactions6-13.However, these microfluidic systems either did not include cultured endothelial cells or were larger than the sizescale relevant to microvascular pathologic processes. A microfluidic platform with cultured endothelial cells that accurately recapitulates the cellular, physical, and hemodynamic environment of the microcirculation is needed to further our understanding of the underlying biophysical pathophysiology of hematologic diseases that involve the microvasculature.
Here, we report a method to create an "endothelialized" in vitro model of the microvasculature, using a simple, single mask microfabrication process in conjunction with standard endothelial cell culture techniques, to study pathologic biophysical microvascular interactions that occur in hematologic disease. This "microvasculature-on-a-chip" provides the researcher with a robust assay that tightly controls biological as well as biophysical conditions and is operated using a standard syringe pump and brightfield/fluorescence microscopy. Parameters such as microcirculatory hemodynamic conditions, endothelial cell type, blood cell type(s) and concentration(s), drug/inhibitory concentration etc., can all be easily controlled. As such, our microsystem provides a method to quantitatively investigate disease processes in which microvascular flow is impaired due to alterations in cell adhesion, aggregation, and deformability, a capability unavailable with existing assays.

A method to culture an endothelial cell monolayer throughout the entire inner 3D surface of a microfluidic device with microvascular-sized channels (&lt;30 &mu;m) is described. This in vitro microvasculature model enables the study of biophysical interactions between blood cells, endothelial cells, and soluble factors in hematologic diseases.

Advances in  microfabrication techniques have enabled the production of inexpensive and  reproducible microfluidic systems for conducting biological and ...

The Submerged Printing of Cells onto a Modified Surface Using a Continuous Flow Microspotter

The printing of cells for microarray applications possesses significant challenges including the problem of maintaining physiologically relevant cell phenotype after printing, poor organization and distribution of desired cells, and the inability to deliver drugs and/or nutrients to targeted areas in the array. Our 3D microfluidic printing technology is uniquely capable of sealing and printing arrays of cells onto submerged surfaces in an automated and multiplexed manner. The design of the microfluidic cell array (MFCA) 3D fluidics enables the printhead tip to be lowered into a liquid-filled well or dish and compressed against a surface to form a seal. The soft silicone tip of the printhead behaves like a gasket and is able to form a reversible seal by applying pressure or backing away. Other cells printing technologies such as pin or ink-jet printers are unable to print in submerged applications. Submerged surface printing is essential to maintain phenotypes of cells and to monitor these cells on a surface without disturbing the material surface characteristics. By printing onto submerged surfaces, cell microarrays are produced that allow for drug screening and cytotoxicity assessment in a multitude of areas including cancer, diabetes, inflammation, infections, and cardiovascular disease.

This 3D microfluidic printing technology prints arrays of cells onto submerged surfaces. We describe how arrays of cells are delivered microfluidically in 3D flow cells onto submerged surfaces. By printing onto submerged surfaces, cell microarrays were produced that allow for drug screening and cytotoxicity assessment in a multitude of areas.

The printing of cells for microarray applications possesses significant challenges including the problem of maintaining physiologically relevant cell ...

Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and other matrix-modifying oxidants/enzymes released during inflammation) are implicated in triggering pathological changes in cellular function, phenotype and viability in a number of diseases. Here, we describe how cell-substrate impedance and live cell imaging approaches can be readily employed to accurately quantify real-time changes in cell adhesion and de-adhesion induced by matrix modification (using endothelial cells and myeloperoxidase as a pathophysiological matrix-modifying stimulus) with high temporal resolution and in a non-invasive manner. The xCELLigence cell-substrate impedance system continuously quantifies the area of cell-matrix adhesion by measuring the electrical impedance at the cell-substrate interface in cells grown on gold microelectrode arrays. Image analysis of time-lapse differential interference contrast movies quantifies changes in the projected area of individual cells over time, representing changes in the area of cell-matrix contact. Both techniques accurately quantify rapid changes to cellular adhesion and de-adhesion processes. Cell-substrate impedance on microelectrode biosensor arrays provides a platform for robust, high-throughput measurements. Live cell imaging analyses provide additional detail regarding the nature and dynamics of the morphological changes quantified by cell-substrate impedance measurements. These complementary approaches provide valuable new insights into how myeloperoxidase-catalyzed oxidative modification of subcellular extracellular matrix components triggers rapid changes in cell adhesion, morphology and signaling in endothelial cells. These approaches are also applicable for studying cellular adhesion dynamics in response to other matrix-modifying stimuli and in related adherent cells (e.g., epithelial cells).

Here, we present a protocol to continuously quantify cell adhesion and de-adhesion processes with high temporal resolution in a non-invasive manner by cell-substrate impedance and live cell imaging analyses. These approaches reveal the dynamics of cell adhesion/de-adhesion processes triggered by matrix modification and their temporal relationship to adhesion-dependent signaling events.

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and ...

Multicolor Fluorescence Detection for Droplet Microfluidics Using Optical Fibers

Fluorescence assays are the most common readouts used in droplet microfluidics due to their bright signals and fast time response. Applications such as multiplex assays, enzyme evolution, and molecular biology enhanced cell sorting require the detection of two or more colors of fluorescence. Standard multicolor detection systems that couple free space lasers to epifluorescence microscopes are bulky, expensive, and difficult to maintain. In this paper, we describe a scheme to perform multicolor detection by exciting discrete regions of a microfluidic channel with lasers coupled to optical fibers. Emitted light is collected by an optical fiber coupled to a single photodetector. Because the excitation occurs at different spatial locations, the identity of emitted light can be encoded as a temporal shift, eliminating the need for more complicated light filtering schemes. The system has been used to detect droplet populations containing four unique combinations of dyes and to detect sub-nanomolar concentrations of fluorescein.

Multicolor fluorescence detection in droplet microfluidics typically involves bulky and complex epifluorescence microscope-based detection systems. Here we describe a compact and modular multicolor detection scheme that utilizes an array of optical fibers to temporally encode multicolor data collected by a single photodetector.

Fluorescence assays are the most common readouts used in droplet microfluidics due to their bright signals and fast time response. Applications such as ...

Training Persons with Spinal Cord Injury to Ambulate Using a Powered Exoskeleton

Powered exoskeletons have become available for overground ambulation in persons with paralyses due to spinal cord injury (SCI) who have intact upper extremity function and are able to maintain upright balance using forearm crutches. To ambulate in an exoskeleton, the user must acquire the ability to maintain balance while standing, sitting and appropriate weight shifting with each step. This can be a challenging task for those with deficits in sensation and proprioception in their lower extremities. This manuscript describes screening criteria and a training program developed at the James J. Peters VA Medical Center, Bronx, NY to teach users the skills needed to utilize these devices in institutional, home or community environments. Before training can begin, potential users are screened for appropriate range of motion of the hip, knee and ankle joints. Persons with SCI are at an increased risk of sustaining lower extremity fractures, even with minimal strain or trauma, therefore a bone mineral density assessment is performed to reduce the risk of fracture. Also, as part of screening, a physical examination is performed in order to identify additional health-related contraindications. 
Once the person has successfully passed all screening requirements, they are cleared to begin the training program. The device is properly adjusted to fit the user. A series of static and dynamic balance tasks are taught and performed by the user before learning to walk. The person is taught to ambulate in various environments ranging from indoor level surfaces to outdoors over uneven or changing surfaces. Once skilled enough to be a candidate for home use with the exoskeleton, the user is then required to designate a companion-walker who will train alongside them. Together, the pair must demonstrate the ability to perform various advanced tasks in order to be permitted to use the exoskeleton in their home/community environment.

Training a person with paralysis to ambulate using a powered exoskeleton may present challenges. The goals are to present the candidate selection criteria and the training procedures for exoskeletal-assisted walking and other mobility skills that can be progressed as the participant's skill level improves.

Powered exoskeletons have become available for overground ambulation in persons with paralyses due to spinal cord injury (SCI) who have intact upper ...

Melt Electrospinning Writing of Three-dimensional Poly(&#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology with great potential for biomedical applications. The technique facilitates the direct deposition of biocompatible polymer fibers to fabricate well-ordered scaffolds in the sub-micron to micro scale range. The establishment of a stable, viscoelastic, polymer jet between a spinneret and a collector is achieved using an applied voltage and can be direct-written. A significant benefit of a typical porous scaffold is a high surface-to-volume ratio which provides increased effective adhesion sites for cell attachment and growth. Controlling the printing process by fine-tuning the system parameters enables high reproducibility in the quality of the printed scaffolds. It also provides a flexible manufacturing platform for users to tailor the morphological structures of the scaffolds to their specific requirements. For this purpose, we present a protocol to obtain different fiber diameters using melt electrospinning writing (MEW) with a guided amendment of the parameters, including flow rate, voltage and collection speed. Furthermore, we demonstrate how to optimize the jet, discuss often experienced technical challenges, explain troubleshooting techniques and showcase a wide range of printable scaffold architectures.

Melt Electrospinning Writing of Three-dimensional Poly(&amp;#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This protocol serves as a comprehensive guideline to fabricate scaffolds via electrospinning with polymer melts in a direct writing mode. We systematically outline the process and define the appropriate parameter settings for achieving targeted scaffold architectures.

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology ...

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for ...