We thank Jean-Baptiste Sibarita and Daniel Choquet (IINS, CNRS/University of Bordeaux) for their kind support with the single-molecule analysis. We especially thank Nick Valmas and Jessica Mcgaw (QBI, University of Queensland) for their help with the video recording, voice-over as well as figure graphic design. This work was supported by the Australian Research Council (AR) discovery project (DP170100125 to F.A.M.), the Australian Research Council (LIEF Grant LE0882864 to F.A.M.), Future Fellowship (FT100100725 to B.V.S.), Australian Research Council (LIEF Grant LE130100078 to F.A.M.) and NHMRC Project grant (APP1103923 to B.V.S.). F.A.M. is a NHMRC Senior Research Fellow (ONT1060075).

1. Design of Transgenic Drosophila Expressing mEos2-Tagged Proteins
<ol>
	<li>Select the protein to be tagged with the mEos2 protein. To increase the imaging success rate, select proteins with a transmembrane domain in the neuronal plasma membrane14 (e.g., Syntaxin-1A), proteins on synaptic vesicles or autophagosomes26,27 (e.g., Synaptobrevin-2), or cytosolic proteins with close interaction with neuronal plasma membrane proteins (e.g., Munc18-1).</li>
	<li>Construct transgenes encoding the mEos2-tagged protein (Figure 1). Design the forward and reverse primers for the protein and mEos2. 
	NOTE: The mEos2 coding sequence can be amplified from the pmEos2-N1 plasmid, which is designed to fuse to the C-terminus of the protein of interest. Cloning can for example be performed by in vivo recombination in Saccharomyces cerevisiae using the pFL44S-attB-MCS-w+ vector14,28, alternately other cloning methods could be used such as Gibson assembly protocol29.
	<ol>
		<li>Linearize the vector with BamHI and XbaI and co-transform into ura3- yeast cells together with the PCR fragments encoding mEos2 and the protein of interest. Inject Drosophila embryos with the construct to generate a transgenic fly line30.</li>
	</ol>
	</li>
	<li>Rear the transgenic fly cultures on standard yeast and molasses medium or some other suitable substitute contained in plastic vials. To ensure healthy and fairly large synaptic boutons31, avoid overcrowding flies in the vials, and flip flies to new vials after each day of egg-laying; this ensures the third instar larvae are well fed and reach proper size by the time they commence wandering on the medium. Raise the transgenic fly at room temperature (25 &#176;C). 
	NOTE: Larger synaptic boutons tend to yield larger quantity of proteins visualized during imaging.</li>
</ol>
2. Dissection of Third Instar Larva
<ol>
	<li>Make a cylindrical or semicylindrical shaped polydimethylsiloxane (PDMS) base &#8804;20 mm in diameter and &#8804;20 mm in height using an appropriate silicone elastomer kit (Figure 2A). Mix silicone elastomer with a silicone hardening agent at a ratio of 10:1.
	<ol>
		<li>Stir the mixture thoroughly for 5 min. Pour the mixture into a mold with the dimsensions listed above. Let it harden in an oven at 100 &#176;C for 45 min. The PDMS base will serve as the foundation on which to carry out dissection. Ensure the PDMS base fits into the well of a glass-bottomed culture dish.</li>
	</ol>
	</li>
	<li>Select a wandering third instar larva from the vial wall. Use an optical stereo-microscope at 4.5 magnification to visualize the larva placed on the PDMS cylindrical base with the dorsal side up.</li>
	<li>Use curved and angled tweezers to stick minutien pins on the head over the mouth hooks, between the two extending anterior spiracles, and on the tail over the anus, between the two posterior spiracles. Add 40 &#181;L of Schneider&#39;s insect medium at room temperature onto the immobilized larva. 
	NOTE: Hemolyph-like solutions (HL3 or HL6) may also be used in the place of Schneider&#39;s solution32,33.</li>
	<li>Provide access through the dorsal side of the abdominal body wall of the larva by use of a minutien pin to incise into the midline of the wall.</li>
	<li>Use spring scissors to cut the body wall open from the incision point. Cut along the midline in the anterior-posterior direction34.</li>
	<li>Use fine forceps and spring scissors to disembowel the larva: remove the internal organs including the salivary glands, trachea and gut. Wash the larva twice with Schneider&#39;s insect medium.</li>
	<li>With fine forceps, hold the two body wall flaps, gently stretch them out, and stick two minutien pins on each side. This should produce a hexagon-shaped preparation (Figure 2A).</li>
	<li>Detach the brain from the ventral nerve cord using spring scissors to decrease the possibility of muscle movement during imaging; this also serves to hold the preparation flat against the imaging dish. Use the curved tweezers to push all six minutien pins into the PDMS base. 
	NOTE: Only a small portion of the pins should be embedded within the abdominal wall.</li>
	<li>To confirm that the larva has survived the dissection procedure, slightly poke the ventral nerve cord, this should elicit a peristaltic contraction of the larva abdominal wall.</li>
</ol>
3. Slightly Oblique Illumination Single-Particle Tracking Microscopy
<ol>
	<li>Invert the dissected larva-PDMS base onto a glass-bottomed culture dish. Fill the imaging dish with 2 mL of room temperature Schneider&#39;s insect medium (Figure 2B).</li>
	<li>Locate motor neurons in the second and third segments with synapses on abdominal muscles 6 and 735 using the 63x/1.2 NA water-immersion objective on a fluorescent total internal reflection fluorescence (TIRF) microscope.</li>
	<li>Use the eyepieces of the microscope to locate the dissected larva; identify muscle 6 by use of bright field illumination.</li>
	<li>Use the microscope acquisition software (see Table of Materials), in TIRF configuration; slide the collimator camera angle to 1&#176; to increase the penetration depth, otherwise move out of TIRF critical angle to achieve oblique illumination.</li>
	<li>Switch the 488 nm laser on to visualize mEos2 in green. Use a low laser power (less than 5% of 22.8 mW) to avoid bleaching the green fluorescence. Locate the neuromuscular junctions (which look like beads on a string) and acquire a low resolution TIRF image.</li>
	<li>Simultaneously switch on the 405 nm laser (photoconverts mEos2 from green to red species) and the 561 nm laser (image the photoconverted red mEos2) to stochastically localize single mEos2 fluorescent proteins. 
	NOTE: Use of low 405 nm laser power is recommended (0.005 to 0.9% of 4.78 mW) as this allows for a relatively constant photoconversion rate during movie acquisition. Use between 50 and 75% of the 561 nm laser (20.1 mW), as this is sufficient to acquire fluorescence from the red mEos2 species without adversely affecting the morphology of the muscle tissue surrounding the neuromuscular junction.</li>
	<li>For each neuromuscular junction chain, acquire a time series composed of 15,000 frames or more at 33 Hz. Save as.czi format in order to preserve the accompanying meta-data for possible reference.</li>
	<li>Use ImageJ to convert &#39;.czi&#39; movie files to &#39;.tiff&#39; files.</li>
	<li>Analyze the acquired movies with suitable single-molecule tracking analytical software11,14,36 such as TrackMate on FIJI/ImageJ37 (see Table of Materials). Locates the &#39;x&#39; and &#39;y&#39; coordinates of each localization by the Gaussian fit of the intensity point spread function (PSF). 
	Note: Troubleshooting: If green fluorescence is not observed or is too dim at the neuromuscular junction upon exposure to the 488 nm laser, briefly switch on the photoconverting and imaging lasers (405 nm and 561 nm), as this helps to produce more of the red species of mEos2. Then switch back to the 488 nm laser and take a TIRF image.</li>
</ol>
4. Single-Molecule Localization in Fixed Larvae
<ol>
	<li>To measure the protein nanocluster size via PALM, replace the Schneider&#39;s insect medium after dissection with a fixative - 4% paraformaldehyde diluted in phosphate buffered saline (PBS).</li>
	<li>Incubate the larva for 30 min at room temperature.</li>
	<li>Remove the dissection pins and place the fixed larva in a 1.7 mL clear snap-lock microcentrifuge tube.</li>
	<li>Dissect and fix as many other larvae as needed, then wash them three times with PBS.</li>
	<li>Replace the PBS with block buffer (a solution of PBS, 0.1% Triton X-100, and 3% normal goat serum).</li>
	<li>Wash the larvae with PBS three times, then incubate with the necessary primary antibody (diluted in the block buffer solution).</li>
	<li>Incubate overnight at 4 oC.</li>
	<li>Wash the larvae three times with PBS, then incubate with the necessary secondary antibody (diluted in the block buffer solution).</li>
	<li>Wash the larvae three times with PBS, attach the larva back onto the PDMS base, and image as previously described for single-molecule tracking.</li>
</ol>

The authors have nothing to disclose.

This protocol describes single-molecule tracking of mEos2-tagged proteins at the neuromuscular junction of the Drosophila third instar larva. To avoid over-expression of transgenic protein, Syntaxin-1A-mEos2 expression was driven by endogenous Syntaxin-1A promoter. Studies of synaptic protein mobility have mostly been limited to in vitro investigations of cultured cells and cortical neurons9,11,12,13. Whether these proteins exhibit similar mobility signatures in a live animal is largely unknown. To visualize single-protein mobility in vivo, limitations such as optical transparency, accessibility, and penetration depth have to be overcome. By dissecting the larval preparation and visualizing the neuromuscular junctions embedded on the surface of the muscle, we avoid the issue of optical transparency and accessibility. Attempts to image single-molecule mobility in normal TIRF configuration proved unsuccessful. However, use of slightly oblique illumination allows for increased excitation laser penetrating depth. Furthermore, the minutien pins used to immobilize the larva helped to avoid any possible adverse effect of anesthetic use on protein mobility. It is possible to use higher magnification objectives, such as the 100x oil immersion objectives, to visualize single molecules in vivo. However, increased magnification may increase the occurance of drifts out of focus. In addition, we have used a lower intensity (~30%) 561 nm laser successfully to image single molecules at motor nerve terminals. Additional testing may be required to use lower laser power.
This protocol is suitable to visualize the mobility of synaptic proteins within the proximity of the neuronal plasma membrane. Using this approach, the mobility of the SNARE protein - Syntaxin-1A and the autophagosome bound protein Atg18a - have been successfully imaged in vivo14,26. The mobility of proteins on motor neuron axons can also be investigated27. However, investigation of the mobility of proteins on the brain or ventral nerve cord may prove difficult to assess due to the high background signal produced by the underlying tissue; in addition, visualizing protein mobility on the same brain structure will require fluorescently tagging the brain structure (to ensure replicability), and use of dual-camera imaging would be needed.
Current methods available for super-resolution microscopy in Drosophila are mostly limited to imaging embryos, rather than the more developed third instar larva38,39. The main issue with these protocols is that they yield a low number of trajectories in the area imaged, and hence prevent in-depth analysis of mobility states22,40. Our in vivo single molecule tracking protocol has the capacity to generate a high number of trajectories, and could therefore be used in other animal models such as Caenorhabiditis elegans and zebrafish. Indeed, every NMJ chain generated ~2,400 trajectories making it suitable for analyzing the different mobile states and transitions between these states14,27. Additionally, many in vivo single molecule imaging strategies rely on the use of anesthetics to immobilize the animals20,22, which might alter the physiologic function under which the imaging is done. Here we optimized an &#39;anesthesia-free&#39; protocol to immobilize the larvae using minutien pins.
A potential use of this protocol may be for visualizing the mobility of proteins in three dimensions. Recent advances in super-resolution microscopy allow protein mobility to be visualized in vitro in &#39;x&#39;, &#39;y&#39;, and &#39;z&#39; dimensions41,42. Our current method does not account for the mobility of proteins in the &#39;z&#39; axis. Yet the Drosophila motor nerve terminal is a spherical structure and a valuable future approach will be to quantify protein mobility in a three-dimensional volume43. Imaging in the z-dimension is feasible using an astigmatic lens. Alternatively, in TIRF mode44,45, the z-resolution is also increased. However, in our experiments, we have used slightly oblique illumination to visualize single molecules of syntaxin1A-mEos2 without the astigmatic lens.
In conclusion, availability of both a transgenic Drosophila fly stock expressing proteins tagged with a photoconvertible protein, a PDMS base to dissect the transgenic larva, as well as a TIRF microscope equipped with the possibility of changing the angle of illumination are the most important tools needed to visualize single proteins as described in this protocol.

Neuronal communication relies on neurotransmitter release, which occurs through the regulated fusion of neurotransmitter-containing synaptic vesicles with the plasma membrane1. This process called exocytosis2,3,4,5,6 is highly dynamic and can be up- or down-regulated according to fluctuations in the rate of stimulation7. The proteins involved in these processes are subject to Brownian motion, and therefore display nanoscopic organization capable of sustaining a range of binding actions that underpin these physiological functions. Plasma membrane proteins are highly dynamic, allowing lateral trapping of molecules into nanoclusters on the plasma membrane7,8,9,10,11,12. As such, investigating the mobility of these proteins helps to elucidate their mode of action8. Synaptic proteins such as the soluble N-ethylmaleimide sensitive factor attachment protein receptors (SNAREs), for example Syntaxin-1A, are now known to exhibit lateral fast mobility, as well as lateral trapping within nanoclusters that may serve as molecular depots and sites for vesicle fusion9,13,14,15.
The recent development of photoconvertible fluorescent proteins, such as monomeric (m)Eos16,17 and Kaede18, has allowed the nanoscopic resolution of neuronal plasma membrane proteins through the use of approaches such as photo-activated localization microscopy (PALM) and stochastic optical reconstruction microscopy (STORM)14,15,19. These developments have enabled investigations into the mobility and nanoclustering of biological proteins in cultured live cells and neurons. More recently, studies have been carried out to elucidate (at similar high resolutions) the dynamics and organization of these membrane proteins in the neurons of live intact organisms such Mus musculus, Caenorhabditis elegans, and Drosophila melanogaster14,20,21,22.
Investigating the dynamics and organization of a protein in vivo requires not only use of the recently developed super-resolution imaging tools (such as PALM, STORM, and photoconvertible fluorophores), but also the ability to overcome hindrances such as accessibility of the protein and the penetration depth of the imaging laser. Imaging intracellular proteins in an intact animal is inherently challenging as is imaging proteins on structures embedded deep within living tissues of the intact animal, such as the hippocampus of an intact mouse. Hence, proteins on or close to the surface of the tissue are more easily imaged. Another difficulty with imaging in vivo is in ensuring reproducibility across animals. As the anatomy might differ from one sample to the other, selecting a structure with ease of replication in different intact animal samples could also prove challenging. The use of stereotypical structures, such as synapses, help with overcoming some of these difficulties.
Recent studies have imaged protein mobility in animals with an optically transparent epidermis such as Caenorhabditis elegans22, while other investigations have imaged protein organization on the surface of a tissue/organ, for example actin imaging in cortical neurons through the skull of a live mouse21. In some cases, anesthetics have been used to keep the animal immobile; however, specific anesthetics may adversely affect the protein of interest. Alternative means to keep animals immobile during in vivo imaging should therefore be explored to minimize error.
Another caveat to super-resolution imaging in vivo is that the lasers used to illuminate the proteins of interest could adversely affect the surrounding tissue, and hence keeping the excitation intensity as low as possible is recommended. Illumination of the surrounding tissue by the laser also tends to increase the background signal. The use of low excitation intensity helps reduce the background, the caveat being it could also lead to a decrease in fluorescent protein photon yield. Background subtraction during analysis could be used as an alternative means to reduce the background signal prior to image analysis.
Drosophila presents an ideal organism for in vivo imaging as it provides well-established genetic tools for the investigation of specific protein function. The upstream activating sequence (UAS)-Gal4 system enables temporal and spatial expression of proteins and can therefore be used to manipulate neurotransmission23, such that thermo-genetic stimulation using Drosophila transient receptor potential sub-family A1 (dTRPA1)24 or opto-genetic stimulation using far-red-shifted CsChrimson25 can be combined with imaging14. We have overcome many of the complications of performing in vivo single-molecule imaging in this system and now describe how single-particle tracking can be carried out in Drosophila melanogaster third instar larvae.

<table border="0" cellpadding="0" cellspacing="0" width="682">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:267px;">PDMS (Sylgard 184 silicone elastomer kit)</td>
			<td style="width:163px;">Dow Corning Corporation</td>
			<td colspan="2" style="width:253px;">000000000001064291</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Glass-bottomed Culture Dish</td>
			<td>In Vitro Scientific</td>
			<td>D29-20-1.5-N</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Optical Stereo-microscope</td>
			<td>Olympus</td>
			<td>SZ51</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Curved Tweezers (Style 5/45)</td>
			<td>Electron Microscopy Sciences</td>
			<td>0208-5AC-PO</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Minutien Pins (0.1mm)</td>
			<td>Fine Science Tools</td>
			<td>26002-10</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Schneider&#39;s Insect Medium</td>
			<td>Sigma, Life Sciences</td>
			<td>S0146</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Vannas Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-00</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Fine Forceps (Dumont #5)</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">ELYRA PS.1 Microscope</td>
			<td>Zeiss</td>
			<td>PS.1</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Phosphate Buffered Saline (PBS)</td>
			<td>Lonza</td>
			<td>17-515Q</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Paraformaldehyde (16%)</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>T-9284</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Normal Goat Serum</td>
			<td>Sigma-Aldrich</td>
			<td>G9023</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Maxyclear snaplock microcentrifuge Tube (1.7 mL)</td>
			<td>Axygen</td>
			<td>MCT-175-C</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Zen Black acquisition software</td>
			<td>Carl Zeiss, 2012</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:267px;">Metamorph software</td>
			<td style="width:163px;">Molecular Devices</td>
			<td style="width:87px;">7.7.8</td>
			<td>PALM Tracer Plugin</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Fly strain</td>
			<td></td>
			<td colspan="2">w1118; HA-Sx1A-mEos2</td>
		</tr>
	</tbody>
</table>

in vivo single-molecule tracking at the drosophila presynaptic motor nerve terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

Using the protocol listed above, Syntaxin-1A-mEos2 transgenic Drosophilamelanogaster were generated (Figure 1). Transgenic third instar Drosophila larvae were dissected on the PDMS cylindrical base (Figure 2A-F). To visualize single molecules of Syntaxin-1A, we used PALM on a TIRF microscope with slightly oblique laser illumination14.
Single molecules of Syntaxin-1A were tracked on the presynaptic motor nerve terminals on muscle 6 of the second abdominal segment. The green fluorescence of mEos2 could be detected as shown by the low-resolution image of Syntaxin-1A-mEos2 on type 1b boutons when excited using the 488 nm laser (Figure 3A). In the imaging experiments, the 405 nm and 561 nm excitation laser imaging depths were 133.5 nm and 165.6 nm, respectively. The green image was acquired as an average of 16 individual frames. This green image was used to create the region of interest used to limit analysis of localizations in the photoconverted movie to the presynaptic motor nerve terminals alone.
The analysis of the acquired photoconverted sptPALM movies generated a super-resolved intensity, diffusion coefficient, and trajectory maps (Figure 3A). Syntaxin-1A-mEos2 mobility was further quantified by analysis of the mean square displacement (MSD) of the individual trajectories (Figure 3B). Statistical comparisons can be done using the area under the MSD curve (Figure 3B). Further analysis of mobility yields the diffusion coefficient distribution of Syntaxin-1A-mEos2, and this can be statistically evaluated by comparison of the mobile to the immobile populations (Figure 3C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56952/56952fig1.jpg" /> 
Figure 1: Generating mEos2-tagged constructs. The pFL44S-attB-MCS-w+ vector is linearized with BamHI and XbaI. Overlapping polymerase chain reaction (PCR) fragments encoding the protein of interest (POI) and mEos2, respectively, can be cloned, for example, by in vivo recombination in ura3- yeast cells or by Gibson assembly. Note that to generate the Syntaxin-1A-mEos2 transgene, we cloned the construct flanked by the endogenous Syntaxin-1A promoter and terminator sequences. The resulting construct can subsequently be injected into Drosophila embryos to create a transgenic fly line expressing the protein of interest (POI) fused to mEos2. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56952/56952fig1large.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/56952/56952fig2.jpg" /> 
Figure 2: Drosophila larva dissection and oblique illumination. (A) Schema showing Drosophila larva dissection carried out on half-cylindrical PDMS base. A filleted larva was held onto the PDMS gel by use of minutien pins. (B-C) The larva-PDMS prep was placed inside a glass-bottomed culture dish containing Schneider&#39;s insect medium. A TIRF microscope in a slightly oblique illumination configuration was used to visualize Syntaxin-1A-mEos2 in the motor nerve terminals. (D-F) A half-cylindrical PDMS base with dissected Drosophila larva. The larva-PDMS preparation was placed in an imaging dish filled with Schneider&#39;s medium, and the larva was imaged on the super-resolution PALM microscope. Scale bar, 10 mm. (This figure has been modified from14). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56952/56952fig2large.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/56952/56952fig3.jpg" /> 
Figure 3: In vivo tracking of Syntaxin-1A-mEos2 at the presynaptic motor nerve terminal. (A) A low-resolution TIRF image of Syntaxin-1A-mEos2 on a type 1b motor nerve terminal. Analysis of photoconversion movies imaged at 33 Hz generated sptPALM super-resolved intensity, diffusion coefficient, and trajectory maps. 5,309 individual trajectories were imaged over 16,000 frames movie acquisition. Scale bar, 3 &#956;m. (B-C) Mean square displacement (MSD) of Syntaxin-1A-mEos2 was plotted from 6 different motor nerve terminals; area under the MSD curve (AUC) was used to quantify the level of mobility. The diffusion coefficient distribution reveals mobile and immobile Syntaxin-1A populations that can be quantified as ratios of each other; an average of ~2,400 trajectories were obtained per motor nerve terminal (n = 3 larvae). Data presented as mean standard error of mean. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56952/56952fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal at JoVE.com

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

in-vivo-single-molecule-tracking-at-drosophila-presynaptic-motor

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.4K Views.  The University of Queensland. Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

Video: In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of primary afferent fibers mediated by GABAergic axo-axonal synapses (primary afferent depolarization). The strength of primary afferent depolarization can be measured by recording of volume-conducted potentials at the dorsal root (dorsal root potentials, DRP). Pathological changes of presynaptic inhibition are crucial in the abnormal central processing of certain pain conditions and in some disorders of motor hyperexcitability. Here, we describe a method of recording DRP in vivo in mice. The preparation of spinal cord dorsal roots in the anesthetized animal and the recording procedure using suction electrodes are explained. This method allows measuring GABAergic DRP and thereby estimating spinal presynaptic inhibition in the living mouse. In combination with transgenic mouse models, DRP recording may serve as a powerful tool to investigate disease-associated spinal pathophysiology. In vivo recording has several advantages compared to ex vivo isolated spinal cord preparations, e.g. the possibility of simultaneous recording or manipulation of supraspinal networks and induction of DRP by stimulation of peripheral nerves.

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

GABAergic presynaptic inhibition is a powerful inhibitory mechanism in the spinal cord important for motor and sensory signal integration in spinal cord networks. Underlying primary afferent depolarization can be measured by recording of dorsal root potentials (DRP). Here we demonstrate a method of in vivo recording of DRP in mice.

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of ...

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

Optical Monitoring of Living Nerve Terminal Labeling in Hair Follicle Lanceolate Endings of the Ex Vivo Mouse Ear Skin

A novel dissection and recording technique is described for optical monitoring staining and de-staining of lanceolate terminals surrounding hair follicles in the skin of the mouse pinna. The preparation is simple and relatively fast, reliably yielding extensive regions of multiple labeled units of living nerve terminals to study uptake and release of styryl pyridinium dyes extensively used in studies of vesicle recycling. Subdividing the preparations before labeling allows test vs. control comparisons in the same ear from a single individual. Helpful tips are given for improving the quality of the preparation, the labeling and the imaging parameters. This new system is suitable for assaying pharmacologically and mechanically-induced uptake and release of these vital dyes in lanceolate terminals in both wild-type and genetically modified animals. Examples of modulatory influences on labeling intensity are given.

Optical Monitoring of Living Nerve Terminal Labeling in Hair Follicle Lanceolate Endings of the Ex Vivo Mouse Ear Skin

Here we describe a novel preparation for imaging live lanceolate sensory terminals of palisade endings that innervate mouse ear skin hair follicles during staining and destaining with styryl pyridinium dyes.

A novel dissection and recording technique is described for optical monitoring staining and de-staining of lanceolate terminals surrounding hair follicles ...

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an excellent model system for such investigations due to its complex behaviors, powerful genetic techniques, and compact nervous system. Laboratory behavioral assays have long been used with Drosophila to simulate properties of the natural environment and study the neural mechanisms underlying the corresponding behaviors (e.g. phototaxis, chemotaxis, sensory learning and memory)1-3. With the recent availability of large collections of transgenic Drosophila lines that label specific neural subsets, behavioral assays have taken on a prominent role to link neurons with behaviors4-11. Versatile and reproducible paradigms, together with the underlying computational routines for data analysis, are indispensable for rapid tests of candidate fly lines with various genotypes. Particularly useful are setups that are flexible in the number of animals tested, duration of experiments and nature of presented stimuli. The assay of choice should also generate reproducible data that is easy to acquire and analyze. Here, we present a detailed description of a system and protocol for assaying behavioral responses of Drosophila flies in a large four-field arena. The setup is used here to assay responses of flies to a single olfactory stimulus; however, the same setup may be modified to test multiple olfactory, visual or optogenetic stimuli, or a combination of these. The olfactometer setup records the activity of fly populations responding to odors, and computational analytical methods are applied to quantify fly behaviors. The collected data are analyzed to get a quick read-out of an experimental run, which is essential for efficient data collection and the optimization of experimental conditions.

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

We describe here a behavioral setup and data analysis method for assaying olfactory responses of up to 100 vinegar flies (Drosophila melanogaster). This system may be used with single or multiple olfactory stimuli, and adaptable for optogenetic activation or silencing of neuronal subsets.

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an ...

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...