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>
	<li>Prepare the mold for PDMS cuboids.
	<ol>
		<li>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).</li>
		<li>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).</li>
		<li>Remove dust from the inner surface of the plate using sticky tape. The mold is ready for use.</li>
	</ol>
	</li>
	<li>Prepare the PDMS mix.
	<ol>
		<li>Mix 7 g of PDMS base and 0.7 g of curing agent (10:1 ratio) thoroughly in a small container.</li>
		<li>Place the container in a vacuum desiccator for at least 15 min to remove air from the mixture.</li>
		<li>Slowly pour about 5.5 g of PDMS mixture onto the mold to reach a 1&#8211;2 mm thickness (Figure 1B).</li>
		<li>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.</li>
		<li>Cure the PDMS on a flat surface in a heat incubator at 65 &#176;C for 2 h.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
3. Mounting larvae for long-term time-lapse imaging
<ol>
	<li>Prepare the top coverslip for mounting.
	<ol>
		<li>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.</li>
		<li>Remove dust from the surface of the PDMS with sticky tape.</li>
		<li>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.</li>
		<li>Apply a small drop (~1.2 &#956;L) 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.</li>
	</ol>
	</li>
	<li>Prepare the larvae for mounting.
	<ol>
		<li>Using a pair of forceps, clean the larvae in water to remove food from the body surface.</li>
		<li>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&#8211;12 drops (160&#8211;240 &#956;L) of isoflurane onto the dry tissue paper using a plastic transferring pipette and close the lid of the large Petri dish.</li>
		<li>Wait 2&#8211;3 min while monitoring the larvae. Take out the larvae from the large Petri dish once their mouth hooks stop moving.</li>
	</ol>
	</li>
	<li>Mount the animals.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Flip the coverslip upside down and repeat step 3.3.4.</li>
		<li>Moisten a small piece of lens paper (15 mm x 30 mm) with 20 &#181;L&#8211;30 &#181;L of water. Place the moistened lens paper at the bottom of the imaging chamber (Figure 1A,D).</li>
		<li>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).</li>
	</ol>
	</li>
</ol>
4. Imaging
<ol>
	<li>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.</li>
</ol>
5. Recovery of imaging chamber and PDMS cuboids
<ol>
	<li>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.</li>
	<li>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.</li>
</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, the spatial and temporal resolution can be adjusted to fit the imaging application at hand (e.g., to track fast subcellular vesicle movement or to monitor slow global changes of neuronal branch patterns). In addition, this method is compatible with larvae at various developmental stages and does not require special equipment. Thus, LarvaSPA can potentially be used by many Drosophila labs to address diverse questions.
Factors affecting the success of the method, including the nature of the PDMS cuboid and the developmental stage of the larvae 
The size of the PDMS cuboid and the depth of the groove need to match the size of the larvae. A PDMS cuboid too wide for the larvae can cover the head and the tail and cause hypoxia. However, a narrow PDMS cuboid may not be effective in preventing the larvae from moving. A too deep groove similarly would not generate enough pressure to restrain the movement of the larvae. A too shallow groove would not hold enough glue to adhere the larvae to the coverslip. Based on our experience, the following dimensions of the PDMS and the groove are recommended for various stages of larvae: 8 mm x 1 mm x 1 mm PDMS cuboids with 1.5 mm x 1 mm x 0.063 mm grooves for second instar larvae (~48 h AEL); 8 mm x 2 mm x 1 mm PDMS cuboids with 2 mm x 2 mm x 0.126 mm grooves for early third instar larvae (~72 h AEL); 8 mm x 2 mm x 1 mm PDMS cuboids with 2 mm x 2 mm x 0.189 mm grooves for late third instar larvae (~96 h AEL or older).
Wandering third instar larvae (at or older than 120 h AEL) move less compared to younger ones and require less moisture to survive once mounted in the chamber. They also have a thicker cuticle and can endure more physical stress. Therefore, experiments using wandering third instar larvae have the highest success rate. In our hands, more than 80% of wandering third instar larvae survived and remained immobile 12 h after mounting. To prevent larvae from pupariating during imaging, we recommend mounting the larvae between 96 h to 120 h AEL. Younger third instar larvae have a greater chance of escape or death during imaging. Typically, at least 2 out of 6 young third instar larvae mounted in the same chamber would survive and remain immobile 10 h after mounting. Our experience with second instar larvae is limited and the survival rate is hard to estimate. Nevertheless, we have successfully imaged second instar larvae for 7 h using this method.
Potential concerns of long-term imaging 
Although LarvaSPA has been very useful for imaging dendrite growth dynamics and degeneration, there are a few potential caveats to consider. First, in our current method, the larvae are deprived of food for the whole duration of imaging, which may lead to starvation-induced responses in many tissues. We have observed formation of granular structures in epidermal cells around 10 h after mounting, which may result from autophagy. Therefore, the results obtained in the first few hours of imaging should be more physiologically relevant. Results over longer time scales require more cautious interpretation. To minimize this concern, our procedure could potentially be adapted to allow for larval feeding. Second, our method may interfere with the rapid growth of younger larvae due to the physical constraint and the lack of nutrient intake. However, this concern may not apply to older animals. For example, wandering third instar larvae can turn into pupae even after 12 h of continuous imaging, making this method ideal for investigations of early metamorphosis. Third, imaging of deeper structures such as the fat body and the gut presents some challenges. A main reason is that deeper tissues often move during imaging and therefore require motion correction in postprocessing. Additionally, mismatches of refractive indices in the light path can cause spherical aberration when imaging deeper tissues. While oil objectives are appropriate for imaging da neurons because their dendrites are within 15 &#956;m from the body surface, water immersion objectives could be better choices to correct refractive-index mismatches for imaging deeper inside the larvae. Fourth, because C4 da neurons can be activated by UV light28, using UV light during the larval mounting can potentially alter C4 da neuron physiology. Although the light intensity we recommend is far below the levels that robustly activate C4da neurons28, results related to C4da neuronal activity need to be cautiously interpreted. The UV glue has been used for imaging a wide variety of biological samples29,30,31 and does not cause obvious toxicity to the larvae in our hands. Lastly, one should consider the proper microscope setup for in vivo live imaging. For example, resonance scanners and spinning disc confocal microscopes are better options for long-term live imaging because they cause less phototoxicity and photobleaching; multi-photon microscopy is more effective for imaging deeper internal structures in the larvae; long-term imaging could be prone to sample or focus drifting, which could potentially be corrected by post-imaging processing.
The most common causes of failure for LarvaSPA and the recommended solutions to address them
<ol>
	<li>Larvae move too much or escape: Two common reasons could contribute to this problem. The first is that the PDMS groove may be too shallow or too deep for the larvae. Trying another PDMS cuboid with a different groove depth may solve the problem. The second reason is that there is too much moisture in the chamber, weakening the UV glue. Reducing the volume of water added to the lens paper in the chamber may help immobilize the larvae. In addition, try to image only the segments covered by the PDMS cuboid, because the head and the tail are free to move.</li>
	<li>Larvae die during imaging: If a larva dies soon after imaging, it is likely that it was exposed to too much anesthetic. Properly anesthetized larvae should wake up after mounting and show motions including mouth hook extension and retraction and dorsal vessel contraction. To solve this problem, less isoflurane could be used to anesthetize larvae. Transfer a larva out of the anesthesia Petri dish immediately after the mouth hook stops moving. If the larva dies about one hour after imaging, it is usually because of dehydration. Remember to place a piece of moistened lens paper in the imaging chamber before sealing. Another common reason of lethality is that the UV glue blocks the spiracles. Try to limit UV glue to the middle segments of the larva and avoid covering the head and the tail. An overly wide PDMS cuboid can also easily cause the UV glue to block the spiracles.</li>
	<li>Folds on the body wall interfere with imaging: To avoid generating folds during mounting, it is important to fully immobilize the larvae during the anesthesia step. When a larva is paralyzed, it is easier to straighten and stretch the body. For animals older than 96 h AEL, gently dragging the tails before curing the UV glue can effectively reduce fold formation. Dragging the tails of young larvae is not recommended because their body walls are fragile.</li>
</ol>

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.

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			<td>193.33161.3</td>
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			<td>Electron Microscopy Sciences</td>
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			<td>Thermo Fisher Scientific</td>
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			<td>UV glue</td>
			<td>Norland products</td>
			<td>#6106, NOA 61</td>
			<td>Refractive Index 1.56</td>
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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 flatten the larval body wall and physically restrict larval movement. Lastly, a small piece of wet lens paper is placed at the bottom of the chamber to provide moisture. This setup can immobilize Drosophila larvae for longer than 10 h for continuous imaging. Most of the animals are alive after 10 h and can be recovered to grow into the pupal stage. The imaging chamber can accommodate up to nine larvae at once. Figure 1D shows six late third instar larvae mounted in the chamber. The trunks of the larvae are fixed while their heads and tails are free to move (Figure 1E and Video S1). This method has been successfully used to image second instar to wandering third instar larvae with continuous high-resolution imaging for up to 15 h. The chamber is designed for imaging using upright microscopes, but the setup also works for inverted microscopes by simply flipping the chamber.
Here we demonstrate the application of LarvaSPA in studying neuronal dendrite dynamics and dendrite degeneration using class IV da (C4 da) neurons as a model (Figure 2, Figure 3, Videos S2&#8211;S4). C4 da neurons are somatosensory nociceptors located on the larval body wall, whose dendrites innervate the larval epidermis1,20,21,22. C4 da dendrites exhibit highly dynamic growth behaviors throughout the larval development, resulting in complete coverage of the body surface or space-filling23. C4 da neurons have also been successfully used to study dendrite degeneration and regeneration after physical injury24,25,26,27.
For imaging dendrite dynamics, larvae ranging from 48 h after egg laying (AEL) at second instar to 120 h AEL at wandering third instar were mounted in the imaging chamber for time-lapse imaging using point-scanning confocal microscopy. Time-lapse movies were taken with a 3 min interval to capture growth behaviors of C4 da dendrites labeled by ppk-CD4-tdTom or UAS-CD4-tdTom driven by ppk-Gal46. Throughout the imaging period (up to 12 h), the high order dendrite branches of C4 da neurons exhibited complex growth behaviors, including extension, retraction, branch formation, and branch elimination (Figure 2A&#8211;2F), indicating the health of the neurons. Our movies also captured homotypic dendro-dendrite repulsions in which dendrite tips retracted after contacting other dendrites (Figure 2F). Overall, these results demonstrate that time-lapse imaging using LarvaSPA is effective for studying neurodevelopment.
To image dendrite degeneration, we used a MaiTai laser to sever primary dendrites near C4 da neuronal cell bodies. The larvae were recovered and mounted in the chamber for imaging starting from 1.5 h after injury (AI) (Figure 3, Video S4). The movies recorded key events during dendrite degeneration, including dendrite swelling, dendrite fragmentation, and clearance of dendrite debris. In the same experiment, the larval fat body was also engineered to secrete Annexin V-GFP (AV-GFP), a sensor that labels externalized phosphatidylserine (PS) on the cell surface27. We observed specific labeling of the degenerating dendrites by AV-GFP (Figure 3), suggesting that PS was exposed on the surface of degenerating dendrites to serve as an eat-me signal for subsequent clearance by phagocytosis27.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60792/60792fig1.jpg" /> 
Figure 1: The imaging chamber for LarvaSPA mounting. (A) Diagrams of the imaging chamber with detailed specifications, showing both the top view and the side view. The light blue shading in the top view indicates the moistened lens paper. The chamber is sealed by a top coverslip and a bottom coverslip. The position of a mounted larva is illustrated. (B) Diagrams of the PDMS mold (the top view) and the mold after filling with the PDMS mixture (side view). Strips in grey indicate two 1-layer, two 2-layer, and two 3-layer tape strips, respectively. The yellow shading in the side view indicates the PDMS mixture in the mold. The dotted lines indicate where to cut the cured PDMS. The side view of the diagram is not drawn to scale. (C) Photographs showing a top view and a side view of a PDMS cuboid. Scale bar = 1 mm. (D) Photographs showing an imaging chamber before mounting, and an imaging chamber with six immobilized late third instar Drosophila larvae mounted dorsal side up. Scale bar = 10 mm. (E) A closer view of a larva in (D). Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60792/60792fig1alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60792/60792fig2.jpg" /> 
Figure 2: Time-lapse imaging of dendrite dynamics. (A-F) Selected frames from time-lapse movies of class IV da neuron dendrites at specific time points after the start of imaging. Images of the larvae were taken 48 h AEL (A and F), 72 h AEL (B), 96 h AEL (C), and 120 h AEL (D). The neurons are labeled by ppk&#62;CD4-tdTom (A, D, E, and F) or ppk &#62;CD4-tdTom (B and C). Blue arrows indicate dendrite retractions compared to the previous time point. Yellow arrows indicate dendrite extensions compared to the previous time point. (E) Dendrite morphology at the end of 12 h of imaging. (F) Consecutive frames with 3 min intervals in a movie illustrating how a dendrite branch in a second instar larva extended (yellow arrows) and subsequently retracted (blue arrows) after it made contact with another dendrite. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60792/60792fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60792/60792fig3.jpg" /> 
Figure 3: Time-lapse imaging of dendrite degeneration and exposure of an eat-me signal. Selected frames from a time-lapse movie of degenerating dendrites of a class IV da neuron after laser injury. Dendrites were labeled by ppk&#62;CD4-tdTom. The eat-me signal PS on degenerating dendrites was detected by Annexin-GFP (AV-GFP), which is secreted by the fat body. Yellow arrows point to the branches showing AV-GFP labeling. Blue arrows point to dendrites undergoing fragmentation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60792/60792fig3alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Video 1" src="/files/ftp_upload/60792/60792video1a.jpg" /> 
Video S1: A third instar larva is immobilized in the notch of a PDMS cuboid. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60792/Movie_S1.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Video 2" src="/files/ftp_upload/60792/60792video2a.jpg" /> 
Video S2: Dendrites extend and retract dynamically in a second instar larva. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60792/Movie_S2.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Video 3" src="/files/ftp_upload/60792/60792video3a.jpg" /> 
Video S3: Dendrites extend and retract dynamically in a wandering third instar larva. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60792/Movie_S3.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Video 4" src="/files/ftp_upload/60792/60792video4a.jpg" /> 
Video S4: Dendrites degenerate and expose phosphatidylserine after laser injury in a third instar larva. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60792/Movie_S4.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>

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

Nanyang Technological University

Research

JoVE Journal

Biology

7.4K Views.  Cornell University. 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.

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

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Actually what I'm gonna do, mount it in here first and then swap out You. So now I'm gonna aspirate try and be as gentle as possible.Okay. A few seconds just holding.Okay.Alright.So the important features of our lifestyle imaging grid are that, so it's an inverted scope, which means the objective, the samples here in the objective is underneath. So the objective is pointing up and the whole scope, or practically the whole, practically the whole scope, including the objectives and the sample are at 37 degrees. So everything in this plastic box is at 37 degrees.And right on top of the stage we have an air and CO2 mix. That is 5%CO2. So the idea is that the sample is actually in a cell, it's like a cell incubator.And so we have, this is our mixer for the air and the CO2 and, and it gets humidified and heated and then piped onto the, the sample on the stage.Okay. And today we're just gonna be doing fluorescence one channel GFP fluorescence. This is the hose that the CO2 and 5%CO2 goes in and it just fits into the stage holder.That's important. And then we can close these, take these down, and then we go to our first position. And what I'm gonna do is go to the first field, which is my control sample, and I'll pick positions based on what I see on the screen.So I just make sure I actually look into the microscope in each new, well, because the focal plane for each of the wells is different, the, the dish is not perfectly flat on the bottom. So you will need to make fine adjustments when you move from well to the well. So right now I'm on the first, well, what I'll do is I'll focus, so I just hit, I just told, I just turned on the light, the transmitted light for the microscope.And I have make sure all the light is going to the eye pieces. And so I just look in, make sure I get a nice clean field. And then what I'll do is turn that light off, turn the fluorescence light on, make sure the cells that I'm looking at look healthy and they're not apoptotic.And, and I turn that off. And what I do next is send, so now I'm gonna start picking positions, which means I have to te send the light path to the camera. This is the camera right here.So I tell the microscope to send all the camera, they don't see anything in the eye pieces. What I'll do is in the software controlling the microscope, I'll choose the right filter set, which for our purposes is psy. And then set an exposure time.And I know these cells very well and I know that they take a 0.2 second exposure time. So I tell, that's what I tell the software to do. And what I'll do is hit focus.This is, and it's basically just alive. The, A light will turn on and you can, oh, I don't know if you can see cells, but the reason this is a good field is because there's, all the cells are, even in terms of their fluorescence, There are no apoptotic cells. Apoptotic cells tend to be very bright and there's no dirt or schmutz in the field.And you can see that they're actually, this is a metaphase cell and that's a metaphase cell, that's a cell that's probably just exited mitosis. And I can just tell that because it looks like those are, those two sets of chromatin are very nicely oriented to each other. Like they just exited an face.So that's a good field. So, and now that I know that the exposure time and the filter set is correct, and this is gonna be software specific, I'll go and I'll take positions and now the software has memorized the X, Y, Z position of that field. And then when I do pick another field, it's, and I'll, so when I'm picking, when I'm picking new fields, but in the same, well, I'll use the, I'll move the i'll, I'll turn the shutter, I'll open the shutter and I'll look to see what I see on the screen and I'll pick fields based on that versus looking in the field.So I don't see any of my tonics in there. But again, even nicely centered, these cells will go through my PS I continue, here's a good example. Here's a good example of what I would avoid that, see how bright that is?That's very bright. That's probably remnants of an apoptotic cell. And the reason I would avoid that is because since it's so bright, when the software auto focuses, it will autofocus just on that and it won't be in the same plane as all these other cells.And so I won't get any information from that field. That looks nice. There's a, this is a pro metaphase right here, That's A metaphase.So, and I'm gonna take three positions per well. So that was three positions and now I'm going to go to my next Well, so you can, so what I will probably do is take images every 10 minutes. So it's a compromise.Lifestyle imaging is always a compromise, right? You have to image, you have to keep the cells healthy, especially if you're interested in mitosis. So you have to minimize their exposure to the fluorescent line.And, but you wanna get information. So you have to take images, you have to, so you have to compromise, you have to make a choice about what you want to capture, what kind of information you wanna capture. For our purposes, mitosis tends to take about an hour.So if we're taking 10 minute intervals, that's pretty good. We see prophase, we see metaphase, we see anaphase, and we can, we can see when there are, there are treatments or drugs or siRNAs that affect those specific cell cycle stages. So for our purposes, that's enough, but we always want more.We always want more positions or finer time and interval or something like that. So it's a compromise. So, and then I'll hit start.And so our software, it does an autofocusing pass on the first pass. So once I hit start, it should start auto autofocusing and you'll see the shutter open, open and close between auto focusing. So I'll hit start.So that's the shutter opening and closing. The Z motor is engaging and it's actually trying to find the plane with the most contrast, which will be the most in focus. You can actually see on the screen going in and out of finding the best plane of focus that might be worrisome.So what it does is it, it drives the Z motor to find the best, the most in focus plane focus, well the most in image. And then it acquires picture there and then it learns that Z position. And then we'll use that Z position for the next 12 passes.And then we'll do the auto focusing pass again. What Kind of experiments do they do? Okay, so our lab is mostly interested in understanding what regulates mitic progression, or at least I'm most interested in understanding that.So with live cell imaging and especially our multi position, long-term imaging scope, you can take, you can film cells for two or three days entering anding mitosis, and you can image multiple rounds of mitosis so you can understand. And then you can put drugs or RNAs or any kind of perturbations on the cells that you would like to study. So you can understand how, how those drugs or, or genes are, are important in regulating mitotic progression.Because it's not fixed cell work and live cell work, you can understand what, when you need to have a perturbation to affect mitosis or what stage of mitosis is affected. So it's really powerful in that respect. What, what are the main technical elements of this experiments?What are the possible problems? Well, the biggest problem besides getting the rig set up, getting the rig set up is not trivial. A lot of people have the rig, but just making sure it works well for your application.The financial element is not, you know, is not small. So there's the setting up of the apparatus and then, and then once you gather the data, there's a real data analysis issue. It takes these movie, if, so, if you're gathering 40 movies each with 30 cells, so you have 40 in 40 fields that you just filmed and each of those fields has 30 cells, then you have to sit down with each one of those movies and look at each one of those cells in that movie.So it takes a really long time to do data analysis. And that's, so it takes a week to analyze data that it took you two days to collect. So there's a, that's a very big obstacle in terms of gathering lots of data.So it's not, while it's high throughput data collection, it's not high throughput analysis. So what would you make to make this experiment really high throughput? So our lab is in the process of developing a program that does the data analysis automatically.And it's a pretty decent program. It's not perfect. The human eye is always better, but it's getting there.So What are the interesting application of this experiment? Maybe other fields you can mention? Besides cell division?You mean for the, oh, for the live cell imaging? Yes.Well, you can do cell motility assays. You can, depending on the, the microscopy that's involved, you can, you know, look at how cells, you could look at tumor development if you really wanted to.But there, you, you're, if as long as you have the right imaging set up, you can, you're pretty much not limited in terms of anything. Our scope happens to be just a wide field and we use 20 by 20 x objective. So it's a pretty low resolution set up, but that's sufficient for what we need to see.So there are, but there definitely aren't, you wouldn't be limited. Anything that has any dynamic kind of experiment, which is biology, it would be, would really benefit from lifestyle imaging. So calcium imaging, although you couldn't do it on our apparatus, that's a live cell.You definitely need live cell imaging for that. Like I said, patient assays, motility assays, those kinds of things would really benefit from live cell imaging. Would you like to tell a few words about your previous experience?How long do you work on these techniques? What do you you work on before? Wow.So in my PhD I didn't work.I did a lot of confocal microscopy, but no lifestyle imaging. And so during my postdoc I was, I helped Dr.King get this apparatus that, that we just used to set up these fragment, get that up and running. And, and so during my postdoc, I had a lot of experience with both widefield with TimeLapse imaging on our scope, which is the widefield and also TimeLapse imaging with confocal microscopy.And I would say that it's really powerful. You get a lot of information, but it's not easy to get working perfectly. So it really requires a lot of attention and a lot of attention to details at every little step of the process.And, and then you still have problems. So it's a tough application, but you can really get, if, if it's what you need to have to, to study your problem, your scientific problem, then it, it's invaluable in that respect, but it's tough. So find someone who knows what they're doing and use them.That's my advice. So good.

Długoterminowe Imaging komórkach ssaków przy dużym polu Mikroskopia

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Serial Enrichment of Spermatogonial Stem and Progenitor Cells (SSCs) in Culture for Derivation of Long-term Adult Mouse SSC Lines

Spermatogonial stem and progenitor cells (SSCs) of the testis represent a classic example of adult mammalian stem cells and preserve fertility for nearly the lifetime of the animal. While the precise mechanisms that govern self-renewal and differentiation in vivo are challenging to study, various systems have been developed previously to propagate murine SSCs in vitro using a combination of specialized culture media and feeder cells1-3.
Most in vitro forays into the biology of SSCs have derived cell lines from neonates, possibly due to the difficulty in obtaining adult cell lines4. However, the testis continues to mature up until ~5 weeks of age in most mouse strains. In the early post-natal period, dramatic changes occur in the architecture of the testis and in the biology of both somatic and spermatogenic cells, including alterations in expression levels of numerous stem cell-related genes. Therefore, neonatally-derived SSC lines may not fully recapitulate the biology of adult SSCs that persist after the adult testis has reached a steady state.
Several factors have hindered the production of adult SSC lines historically. First, the proportion of functional stem cells may decrease during adulthood, either due to intrinsic or extrinsic factors5,6. Furthermore, as with other adult stem cells, it has been difficult to enrich SSCs sufficiently from total adult testicular cells without using a combination of immunoselection or other sorting strategies7. Commonly employed strategies include the use of cryptorchid mice as a source of donor cells due to a higher ratio of stem cells to other cell types8. Based on the hypothesis that removal of somatic cells from the initial culture disrupts interactions with the stem cell niche that are essential for SSC survival, we previously developed methods to derive adult lines that do not require immunoselection or cryptorchid donors but rather employ serial enrichment of SSCs in culture, referred to hereafter as SESC2,3.
The method described below entails a simple procedure for deriving adult SSC lines by dissociating adult donor seminiferous tubules, followed by plating of cells on feeders comprised of a testicular stromal cell line (JK1)3. Through serial passaging, strongly adherent, contaminating non-germ cells are depleted from the culture with concomitant enrichment of SSCs. Cultures produced in this manner contain a mixture of spermatogonia at different stages of differentiation, which contain SSCs, based on long-term self renewal capability. The crux of the SESC method is that it enables SSCs to make the difficult transition from self-renewal in vivo to long-term self-renewal in vitro in a radically different microenvironment, produces long-term SSC lines, free of contaminating somatic cells, and thereby enables subsequent experimental manipulation of SSCs.

Serial Enrichment of Spermatogonial Stem and Progenitor Cells SSCs in Culture for Derivation of Long-term Adult Mouse SSC Lines

A simple method to derive and maintain spermatogonial stem and progenitor cell lines from adult mice is presented here. The method utilizes feeder cells originating from the somatic cell compartment of the adult mouse testis. This technique is applicable to common mouse strains, including transgenic, knock-out, and knock-in mice.

Spermatogonial stem and progenitor cells (SSCs) of the testis represent a classic example of adult mammalian stem cells and preserve fertility for nearly ...

Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is particularly suited for long-term time-lapse imaging over many hours up to several days. However, an appropriate sample mounting strategy is needed that offers both confinement and normal development of the sample. Multilayer mounting, a new embedding technique using low-concentration agarose in optically clear tubes, now overcomes this limitation and unleashes the full potential of light sheet microscopy for real-time developmental biology.

The development of zebrafish can be followed over days with light sheet microscopy when embryos are embedded in optically clear polymer tubes with low-concentration agarose.

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is ...

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time points. Time intervals can be set through a computer interface that controls the microscope-integrated camera. This kind of microscopy requires both the ability to acquire very rapid events and the signal generated by the observed cellular structure during these events. After the images have been collected, a movie of the entire experiment is assembled to show the dynamic of the molecular events of interest. Time-lapse video microscopy has a broad range of applications in the biomedical research field and is a powerful and unique tool for following the dynamics of the cellular events in real time. Through this technique, we can assess cellular events such as migration, division, signal transduction, growth, and death. Moreover, using fluorescent molecular probes we are able to mark specific molecules, such as DNA, RNA or proteins and follow them through their molecular pathways and functions. Time-lapse video microscopy has multiple advantages, the major one being the ability to collect data at the single-cell level, that make it a unique technology for investigation in the field of cell biology. However, time-lapse video microscopy has limitations that can interfere with the acquisition of high quality images. Images can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. Herein, we describe a protocol for the dynamic acquisition of a specific protein, Parkin, fused with the enhanced yellow fluorescent protein (EYFP) in order to track the selective removal of damaged mitochondria, using a time-lapse video microscopy approach.

Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is regulated spatially and temporally. The use of a fluorescent gene under the control of an immunity-related promoter in combination with an automated fluorescence microscopy is a simple way to understand spatiotemporal regulation of plant immunity. In contrast to the root tissues that have been used for a number of various intravital fluorescent imaging experiments, there exist few fluorescent live-imaging examples for the leaf tissues that encounter an array of airborne microbial infections. Therefore, we developed a simple method to mount leaves of Arabidopsis thaliana plants for live-cell imaging over an extended period of time. We used transgenic Arabidopsis plants expressing the yellow fluorescent protein (YFP) genefused to the nuclear localization signal (NLS) under the control of the promoter of a defense-related marker gene, Pathogenesis-Related 1 (PR1). We infiltrated a transgenic leaf with Pseudomonas syringae pv. tomato DC3000 (avrRpt2) strain (Pst_a2) and performed in vivo time-lapse imaging of the YFP signal for a total of 40 h using an automated fluorescence stereomicroscope. This method can be utilized not only for studies on plant immune responses but also for analyses of various developmental events and environmental responses occurring in leaf tissues.

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

We report a simple and versatile method for performing fluorescent live-imaging of Arabidopsis thaliana leaves over an extended period of time. We use a transgenic Arabidopsis plant expressing a fluorescent reporter gene under the control of an immunity-related promoter as an example for gaining spatiotemporal understanding of plant immune responses.

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is ...

Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and several essential mitochondrial polypeptides. Mitochondrial nucleoids divide and distribute within the dynamic mitochondrial network that undergoes fission/fusion and other morphological changes. High resolution live fluorescence microscopy is a straightforward technique to characterize a nucleoid&#39;s position and motion. For this technique, nucleoids are commonly labeled through fluorescent tags of their protein components, namely transcription factor a (TFAM). However, this strategy needs overexpression of a fluorescent protein-tagged construct, which may cause artifacts (reported for TFAM), and is not feasible in many cases. Organic DNA-binding dyes do not have these disadvantages. However, they always show staining of both nuclear and mitochondrial DNAs, thus lacking specificity to mitochondrial nucleoids. By taking into account the physico-chemical properties of such dyes, we selected a nucleic acid gel stain (SYBR Gold) and achieved preferential labeling of mitochondrial nucleoids in live cells. Properties of the dye, particularly its high brightness upon binding to DNA, permit subsequent quantification of mitochondrial nucleoid motion using time series of super-resolution structured illumination images.

The protocol describes specific labeling of mitochondrial nucleoids with a commercially available DNA gel stain, acquisition of time lapse series of live labeled cells by super-resolution structured illumination microscopy (SR-SIM), and automatic tracking of nucleoid motion.

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and ...