We thank Lingfeng Tang for establishing an earlier version of the LarvaSPA method; Glenn Swan at Cornell Olin Hall Machine shop for making earlier prototypes of the imaging chamber; Philipp Isermann for constructing metal frames and providing suggestions on making PDMS cuboids; Cornell BRC Imaging facility for access to microscopes (funded by NIH grant S10OD018516); Maria Sapar for critical reading of the manuscript. This work was supported by a Cornell Fellowship awarded to H.J.; a Cornell start-up fund and NIH grants (R01NS099125 and R21OD023824) awarded to C.H. H.J. and C.H. conceived the project and designed the experiments. H.J. conducted the experiments. H.J and C.H. wrote the manuscript.

1. Making the imaging chamber
<ol>
	<li>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.</li>
	<li>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.</li>
</ol>
2. Making PDMS cuboids<...

<ol>
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The authors declare no competing interests.

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...

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 structures1,2. 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 microscopy7,8,9. 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) &#34;larva chip&#34;, 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 junctions11,12,13,14,15,16. 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 studied17,18. 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 &#34;Larva Stabilization by Partial Attachment&#34; (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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td>6061 Aluminum bars</td>
			<td>McMaster-Carr</td>
			<td>9246K421</td>
			<td></td>
		</tr>
		<tr>
			<td>3M double-sided tape</td>
			<td>Ted Pella, Inc.</td>
			<td>16093</td>
			<td></td>
		</tr>
		<tr>
			<td>3M Scotch Packaging tape</td>
			<td>3M</td>
			<td></td>
			<td>1.88&#34;W x 22.2 Yards</td>
		</tr>
		<tr>
			<td>DUMONT #3 Forceps</td>
			<td>Fisher Scientific</td>
			<td>50-241-34</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslip</td>
			<td>Azer Scientific</td>
			<td>1152250</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Midwest Veterinary Supply</td>
			<td>193.33161.3</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica Confocal Microscope</td>
			<td>Leica</td>
			<td>SP8 equipped with a resonant scanner</td>
			<td></td>
		</tr>
		<tr>
			<td>Lens paper</td>
			<td>Berkshire</td>
			<td>LN90.0406.24</td>
			<td></td>
		</tr>
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			<td>Petri dishes (medium)</td>
			<td>VWR</td>
			<td>25373-085</td>
			<td></td>
		</tr>
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			<td>Petri dishes (small)</td>
			<td>VWR</td>
			<td>10799-192</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor blade</td>
			<td>Ted Pella, Inc.</td>
			<td>121-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Rectangular petri dish</td>
			<td>VWR</td>
			<td>25384-322</td>
			<td></td>
		</tr>
		<tr>
			<td>SYLGARD 184 kit (PBMS kit)</td>
			<td>Electron Microscopy Sciences</td>
			<td>24236-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Transferring pipette</td>
			<td>Thermo Fisher Scientific</td>
			<td>1371126</td>
			<td></td>
		</tr>
		<tr>
			<td>UV glue</td>
			<td>Norland products</td>
			<td>#6106, NOA 61</td>
			<td>Refractive Index 1.56</td>
		</tr>
		<tr>
			<td>UV lamp (Workstar 2003)</td>
			<td>Maxxeon</td>
			<td>MXN02003</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum desiccator</td>
			<td>Electron Microscopy Sciences</td>
			<td>71232</td>
			<td></td>
		</tr>
		<tr>
			<td>Wipes</td>
			<td>Kimberly-Clark</td>
			<td>Kimwipes</td>
			<td></td>
		</tr>
	</tbody>
</table>

larvaspa, a method for mounting drosophila larva for long-term time-lapse imaging

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.

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 ...

Watch this Scientific Journal Video about LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging at JoVE.com

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

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.

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

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because ...

LarvaSPA is the first method that allows for continuous live imaging of intact drosophila larvae for more than 10 hours with high temporal and spatial resolution. This method is used for revealing dynamic cellular processes of larvae peripheral sensory neurons and for studying many other cellular processes that happen near the larva body wall. This method is easy to use and has less limitation on larva size.Six to nine larvae can be mounted in the imaging chamber at the same time, which makes the experiment possible. The imaging chamber at the PDMS cuboids can be manufactured at a minimal cost and are reusable. This method is particularly useful for studying mechanisms of dendrite development and dendrite degeneration using drosophila larva dendritic authorization neurons.Being able to image the same neuron for hours can reveal how long term global changes of dendritic patterns arise from short term behaviors at the individual branch level. Using PDMS cuboids that match the sizes of larvae will increase the success rate of immobilizing larvae. Please find more useful tapes in our troubleshooting section.After constructing an aluminum block from a machine shop, seal the bottom of the metal frame using a long cover slip with UV glue. Cure the UV clue using a handheld UV lamp for 30 seconds. Attach layers of packaging tape to the inner surface of a rectangular Petri dish or round cell culture plate.Use one layer for second in-star larvae, two layers for early third in-star larvae, or three layers for late third in-star larvae. Cut the tape into trips of specific width with a razor blade, 1.5 millimeters for the one layer tape and two millimeters for the two layer or three layer tape. Leave at least a five millimeter space between the two strips.Remove the tape layers covering the space. Remove dust from the inner surface of the plate using sticky tape. After that, the mold is ready for use.To prepare the PDMS mix, mix seven grams PDMS base in 0.7 grams of curing agent thoroughly in a small container. Place the container in a vacuum desiccator for at least 15 minutes to remove air from the mixture. Slowly pour about 5.5 grams PDMS mixture onto the mold to reach a one to two millimeter thickness.Place the PDMS mixture in the vacuum desiccator again for at least 15 minutes to remove remaining air bubbles from the mixture. Break the last few bubbles with a pipette tip. Cure the PDMS on a flat surface in a heat incubator at 65 degrees Celsius for two hours.Next, 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. For early and late third in-star larvae, cut the PDMS into eight by two by one millimeter cuboids by positioning the groove created by the tape strip at the center of the long side of the cuboid.For second in-star larvae, cut the cuboid to eight by one by one millimeter. To prepare the top cover slip for mounting, choose six PDMS cuboids with grooves matching the sizes of the larvae. Remove dust from the surface of the PDMS with sticky tape.Attach four pieces of double sided tape at the size of 12 by five millimeters on a long cover slip of 22 by 50 millimeters for fixing PDMS cuboids later. Adjust the spaces between the two pieces of double sided tape to the same as the width of the PDMS groove. Apply a small drop 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 tapes on the cover slip.To prepare the larvae for mounting, using a pair of forceps, clean the larvae in water to remove food from the body surface. Place the clean larvae on a small piece of moistened tissue paper in a small Petri dish without a lid and place the small Petri dish into a large Petri dish containing a piece of dry tissue paper. In a chemical hood, use a plastic transferring pipette to apply 160 to 240 microliters of isoflurane onto the dry tissue paper and close the lid of the large Petri dish.Wait two to three minutes while monitoring the larvae. Once their mouth hooks stop moving, take out the larvae from the large Petri dish. Place the immobilized larvae onto the UV glue between the double sided tapes on the cover slip with the dorsal cuticle facing the cover slip.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.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. Wear safety glasses and use a handheld UV lamp to cure the UV glue for four minutes on the high setting.Flip the cover slip upside down and use the UV lamp to cure the UV glue for another four minutes. Place a small piece of lens paper with the size of 15 by 30 millimeters at the bottom of the imaging chamber. Moisten the paper with 20 to 30 microliters of water.Place the cover slip on the chamber so that the larvae are facing the inside of the chamber. Use the UV glue to adhere both ends of the cover slip to the metal surface. The dorsal side of the larvae is ready for imaging under a confocal microscope.After imaging, remove the oil on the top cover slip using lens paper. Detach the top cover slip from the metal frame by cutting into the space between the cover slip and the metal frame with a razor blade. The imaging chamber is ready for reuse.Detach the PDMS cuboids from the top cover slip with forceps. Roll the PDMS cuboids on sticky tape to remove glue residue and dust. The PDMS cuboids are ready for reuse.In this protocol, drosophila larvae were immobilized for longer than 10 hours for continuous imaging. This figure shows six late third in-star larvae mounted in the chamber. The trunks of the larvae were fixed while their heads and tails were free to move.Here, we demonstrate the application of LarvaSPA in studying neuronal dendrite dynamics and dendrite degeneration using class four DA neurons as a model. To successfully immobilize larvae using this method, it is critical to stop exposing the larvae to isoflurane once the mouth hooks stop moving and cure the UV glue before larvae fully wake up. To study dendrite degeneration and regeneration, laser injury can be performed when larvae are immobilized in the imaging chamber.LarvaSPA has helped us to understand how dynamic growing dendrites fail to innervate epidermal cells like in hepatic cell proteoglycans and how phosphatidylserine is exposed on degenerating dendrites after laser injury. Protect eyes with safety glasses while using the UV lamp. Knocking out larvae using isoflurane should be performed in a fume hood.

larvaspa-method-for-mounting-drosophila-larva-for-long-term-time

Research

JoVE Journal

Biology

7.3K visualizações.  Cornell University. LarvaSPA é o primeiro método que permite imagens vivas contínuas de larvas de drosophila intactas por mais de 10 horas com alta resolução temporal e espacial. Este método é usado para revelar processos celulares dinâmicos de neurônios sensoriais periféricos larvas e para estudar muitos outros processos celulares que acontecem perto da parede corporal da larva. Este método é fácil de usar e tem menor limitação no tamanho da larva.Seis a nove larvas podem ser montadas na c...