This work was supported by Natural Science Foundation of China Grant 32070811, and Southeast University (China) Analysis Test Fund 11240090971. We thank the Laboratory of Electron Microscopy and Center of Morphological Analysis, School of Medicine, Southeast University, Nanjing, China.

NOTE: The transmission electron microscopy sample preparation method used in this article has been reported previously16. It is important to note that the selection of reagents and the adjustment of dosage are necessary depending on the sample. There are many toxic chemical reagents used in the sample preparation process, therefore, the operator needs to take certain protective measures, such as wearing protective clothing and gloves and operating in a fume hood.
1. Dissection, fixation and placement procedure
<ol>
	<li>Dissection
	<ol>
		<li>Dissect the wandering late-3rd-instar larvae in Jan solution and obtain the entire body wall based on the standard procedures17. Prepare Jan&#39;s solution as following: 128 mM NaCl, 2 mM KCl, 4 mM MgCl2, 35mM sucrose, 5mM HEPES, pH 7.4 
		NOTE: The body wall of Drosophila larvae refers to the outer covering or integument of the larval stage of the fruit fly Drosophila. This includes the cuticle, epidermal cells, and underlying tissues that form the larval exoskeleton. The body wall provides structural support and protection and serves as a barrier between the larval body and the external environment17.</li>
	</ol>
	</li>
	<li>Fixation
	<ol>
		<li>Fix the larval body wall in the dissection chamber at 4 &#176;C overnight with the fixative. To prepare the fixative, mix 2% glutaraldehyde and 2% formaldehyde in 0.1 M sodium cacodylate buffer (pH 7.4)</li>
		<li>Rinse the samples several times with sodium cacodylate buffer.</li>
		<li>Prepare 1% OsO4 in cacodylate buffer. Fix the samples in 1% OsO4 for 2 h, followed by several rinses in distilled water. 
		NOTE: Dual fixation with aldehydes and OsO4 provides better protection of cellular ultrastructure. Besides, OsO4 can stain the sample tissue, allowing for the contrast between the light and the sample to be more apparent18.</li>
		<li>Stain the Drosophila neuromuscular junction samples with 2% saturated uranyl acetate for 2 h, followed by several rinses with deionized water (Figure 1A).</li>
	</ol>
	</li>
	<li>Sample placement
	<ol>
		<li>Cut two round pieces of metal mesh of similar size (pore size 270 &#181;m) with scissors.</li>
		<li>Place a metal mesh (270 &#181;m) at the base of a flat-bottomed bottle in advance.</li>
		<li>Place the fixed larva samples on the mesh and then place another mesh on them so that the samples are placed between the two meshes.</li>
	</ol>
	</li>
</ol>
2. Dehydration
NOTE In this paper, the ethanol gradient dehydration method was used.
<ol>
	<li>Dehydrate the samples in an increasing concentration gradient of ethanol sequentially (50%, 70%, 85%, 95% once each, 100% twice), each for 20 min at 4 &#176;C.</li>
	<li>Rinse the samples twice with propylene oxide at room temperature for 5 min each time (Figure 1C).</li>
</ol>
3. Infiltration
NOTE: Infiltration is one of the necessary steps for the TEM sample preparation. Infiltration allows for the gradual replacement of dehydrating medium with the embedding medium in the sample to allow filling of the cell interstice with the embedding medium19.
<ol>
	<li>Place the samples in the mixed solution of propylene oxide and epoxy resin at a ratio of 1:1 and 1:2 for 1 h each, followed by pure epoxy resin for 2 h at room temperature.</li>
</ol>
4. Embedding
<ol>
	<li>Prepare a polyethylene film and cut it into a big square (approximately 5 &#215; 5 cm) and a small hollow rectangular support (approximately 2 &#215; 2 cm). Glue the small square with the big square.</li>
	<li>Pre-position sufficient epoxy resin at the center of the big square.</li>
	<li>Place the samples in the pre-positioned epoxy resin with the muscle side facing upwards. Use a stereoscope.</li>
	<li>Add several drops of resin to the samples and then cover them with PE film (a little larger than the spacer) (Figure 1D-E).</li>
	<li>Polymerize the samples at 37 &#176;C, 42 &#176;C and 60 &#176;C for 24 h respectively (Figure 1F).</li>
</ol>
5. Sectioning procedure
<ol>
	<li>Remove the excess parts of the larval body wall with a sharp blade and retain a trapezoid, including the A2/A3 segment. 
	NOTE: The top line should be near the 13th muscle, and the baseline should be away from the sample to the epoxy resin. The height between the topline and baseline is approximately 5-8 mm.</li>
	<li>Remove the trapezoid from the remainder of the sample and adhere it to the resin-filled capsule with the A/B glue.</li>
	<li>Confirm the position of the 6th and 7th muscles of A2 and A3 segments under the light microscopy (Figure 1G) and maintain the tangent plane parallel to the 6th muscle.</li>
	<li>Cut away one-fifth of the sample along the two sides of the A3 segment. 
	NOTE: Remove most of the 6th muscle, and retain 1/3 of the width of the 7th muscle.</li>
	<li>Using an ultrathin microtome, prepare an ultrathin section of the sample starting with the 6th muscle. 
	NOTE: Each slice is approximately 90 nm thick.</li>
	<li>Attached as many slices as possible to each copper mesh.</li>
	<li>Perform transmission electron microscopy as described previously16.</li>
</ol>

<ol>
	<li>Acevedo, N. C., Marangoni, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanostructured+fat+crystal+systems.">Nanostructured fat crystal systems.</a> Annual Review of Food Science and Technology. 6, 71-96 (2015).</li><li>Schorb, M., Haberbosch, I., Hagen, W. J. H., Schwab, Y., Mastronarde, D. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Software+tools+for+automated+transmission+electron+microscopy.">Software tools for automated transmission electron microscopy.</a> Nature Methods. 16 (6), 471-477 (2019).</li><li>Winans, N. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+genomic+investigation+of+ecological+differentiation+between+free-living+and+Drosophila-associated+bacteria.">A genomic investigation of ecological differentiation between free-living and Drosophila-associated bacteria.</a> Molecular Ecology. 26 (17), 4536-4550 (2017).</li><li>Phelps, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstruction+of+motor+control+circuits+in+adult+Drosophila+using+automated+transmission+electron+microscopy.">Reconstruction of motor control circuits in adult Drosophila using automated transmission electron microscopy.</a> Cell. 184 (3), 759-774 (2021).</li><li>Cardona, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+integrated+micro-+and+macroarchitectural+analysis+of+the+Drosophila+brain+by+computer-assisted+serial+section+electron+microscopy.">An integrated micro- and macroarchitectural analysis of the Drosophila brain by computer-assisted serial section electron microscopy.</a> PLoS Biology. 8 (10), (2010).</li><li>Belalcazar, H. M., Hendricks, E. L., Zamurrad, S., Liebl, F. L. W., Secombe, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+histone+demethylase+KDM5+is+required+for+synaptic+structure+and+function+at+the+Drosophila+neuromuscular+junction.">The histone demethylase KDM5 is required for synaptic structure and function at the Drosophila neuromuscular junction.</a> Cell Reports. 34 (7), 108753 (2021).</li><li>Banerjee, S., Venkatesan, A., Bhat, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurexin,+neuroligin+and+wishful+thinking+coordinate+synaptic+cytoarchitecture+and+growth+at+neuromuscular+junctions.">Neurexin, neuroligin and wishful thinking coordinate synaptic cytoarchitecture and growth at neuromuscular junctions.</a> Molecular Cell and Neurosciences. 78, 9-24 (2017).</li><li>Guangming, G., Junhua, G., Chenchen, Z., Yang, M., Wei, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurexin+and+neuroligins+maintain+the+balance+of+ghost+and+satellite+boutons+at+the+Drosophila+neuromuscular+junction.">Neurexin and neuroligins maintain the balance of ghost and satellite boutons at the Drosophila neuromuscular junction.</a> Frontiers in Neuroanatomy. 14, 19 (2020).</li><li>Jia, X. X., Gorczyca, M., Budnik, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrastructure+of+neuromuscular+junctions+in+Drosophila:+comparison+of+wild+type+and+mutants+with+increased+excitability.">Ultrastructure of neuromuscular junctions in Drosophila: comparison of wild type and mutants with increased excitability.</a> Journal of Neurobiology. 24 (8), 1025-1044 (1993).</li><li>Ashley, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retrovirus-like+gag+protein+Arc1+binds+RNA+and+traffics+across+synaptic+boutons.">Retrovirus-like gag protein Arc1 binds RNA and traffics across synaptic boutons.</a> Cell. 172 (1-2), 262-274 (2018).</li><li>Eaton, B. A., Fetter, R. D., Davis, G. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynactin+is+necessary+for+synapse+stabilization.">Dynactin is necessary for synapse stabilization.</a> Neuron. 34 (5), 729-741 (2002).</li><li>Featherstone, D. E., Rushton, E., Broadie, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+regulation+of+glutamate+receptor+field+size+by+nonvesicular+glutamate+release.">Developmental regulation of glutamate receptor field size by nonvesicular glutamate release.</a> Nature Neurosciences. 5 (2), 141-146 (2002).</li><li>Wagner, N., Laugks, U., Heckmann, M., Asan, E., Neuser, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aging+Drosophila+melanogaster+display+altered+pre-+and+postsynaptic+ultrastructure+at+adult+neuromuscular+junctions.">Aging Drosophila melanogaster display altered pre- and postsynaptic ultrastructure at adult neuromuscular junctions.</a> Journal of Comparative Neurology. 523 (16), 2457-2475 (2015).</li><li>Atwood, H. L., Govind, C. K., Wu, C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+ultrastructure+of+synaptic+terminals+on+ventral+longitudinal+abdominal+muscles+in+Drosophila+larvae.">Differential ultrastructure of synaptic terminals on ventral longitudinal abdominal muscles in Drosophila larvae.</a> Journal of Neurobiology. 24 (8), 1008-1024 (1993).</li><li>McCabe, B. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+BMP+homolog+Gbb+provides+a+retrograde+signal+that+regulates+synaptic+growth+at+the+Drosophila+neuromuscular+junction.">The BMP homolog Gbb provides a retrograde signal that regulates synaptic growth at the Drosophila neuromuscular junction.</a> Neuron. 39 (2), 241-254 (2003).</li><li>Guangming, G., Mei, C., Chenchen, Z., Wei, X., Junhua, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+analysis+method+of+neuromuscular+junction+in+Drosophila.+larvae+by+transmission+electron+microscopy.">Improved analysis method of neuromuscular junction in Drosophila. larvae by transmission electron microscopy.</a> Anatomical Science International. 97 (1), 147-154 (2022).</li><li>Brent, J. R., Werner, K. M., McCabe, B. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+larval+NMJ+dissection.">Drosophila larval NMJ dissection.</a> Journal of Visualized Experiments. (24), e1107 (2009).</li><li>Lovri&#263;, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multimodal+imaging+of+chemically+fixed+cells+in+preparation+for+NanoSIMS.">Multimodal imaging of chemically fixed cells in preparation for NanoSIMS.</a> Analytical Chemistry. 88 (17), 8841-8848 (2016).</li><li>Woods, P. S., Ledbetter, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+organelles+at+uncoated+cryofractured+surfaces+as+viewed+with+the+scanning+electron+microscope.">Cell organelles at uncoated cryofractured surfaces as viewed with the scanning electron microscope.</a> Journal of Cell Sciences. 21 (1), 47-58 (1976).</li><li>Jan, L. Y., Jan, Y. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Properties+of+the+larval+neuromuscular+junction+in+Drosophila+melanogaster.">Properties of the larval neuromuscular junction in Drosophila melanogaster.</a> Journal of Physiology. 262 (1), 189-214 (1976).</li><li>Prokop, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+of+the+efferent+system+and+structure+of+neuromuscular+junctions+in+Drosophila.">Organization of the efferent system and structure of neuromuscular junctions in Drosophila.</a> International Review in Neurobiology. 75, 71-90 (2006).</li><li>Lloyd, T. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+p150(Glued)+CAP-Gly+domain+regulates+initiation+of+retrograde+transport+at+synaptic+termini.">The p150(Glued) CAP-Gly domain regulates initiation of retrograde transport at synaptic termini.</a> Neuron. 74 (2), 344-360 (2012).</li><li>Budnik, V., Zhong, Y., Wu, C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+plasticity+of+motor+axons+in+Drosophila+mutants+with+altered+excitability.">Morphological plasticity of motor axons in Drosophila mutants with altered excitability.</a> Journal of Neuroscience. 10 (11), 3754-3768 (1990).</li><li>Zhang, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuroligin+4+regulates+synaptic+growth+via+the+bone+morphogenetic+protein+(BMP)+signaling+pathway+at+the+Drosophila+neuromuscular+junction.">Neuroligin 4 regulates synaptic growth via the bone morphogenetic protein (BMP) signaling pathway at the Drosophila neuromuscular junction.</a> Journal of Biological Chemistry. 292 (44), 17991-18005 (2017).</li><li>Wu, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+pre-synaptic+function+of+shank+protein+in+Drosophila.">A pre-synaptic function of shank protein in Drosophila.</a> Journal of Neuroscience. 37 (48), 11592-11604 (2017).</li><li>Gan, G., Lv, H., Xie, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+identification+and+development+of+neurite+in+Drosophila+ventral+nerve+cord+neuropil.">Morphological identification and development of neurite in Drosophila ventral nerve cord neuropil.</a> PLoS One. 9 (8), 105497 (2014).</li></ol>

The authors have nothing to disclose.

Drosophila larva samples tend to curl up during dehydration, as the samples are thin, making it difficult to accurately locate the neuromuscular junction, thereby increasing the difficulty and workload for the sample preparation. The traditional improvement is to shorten the sample7, but the samples were still curled to different degrees.
In our method, there are two critical steps: first, the samples remain flat throughout the dehydration process due to the re...

An erratum was issued for:&#160;<a href="https://jove.unicartagenaproxy.elogim.com/t/64934">Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae</a>. The Authors section was updated from:
Gan Guangming1,2 
Sheng Qingyuan3 
Ou Yutong1 
Chen Mei1 
Zhang Chenchen1 
1The School of Medicine, Southeast University, 
2The Key Laboratory of Developmental Genes and Human Disease, Southeast University, 
3The School of Life Science and Technology, Southeast University 
Corresponding Authors:&#160;Gan Guangming
to:
Gan Guangming1,2 
Sheng Qingyuan3 
Ou Yutong1 
Chen Mei3 
Zhang Chenchen1 
1The School of Medicine, Southeast University, 
2The Key Laboratory of Developmental Genes and Human Disease, Southeast University, 
3The School of Life Science and Technology, Southeast University 
Corresponding Authors:&#160;Gan Guangming,&#160;Sheng Qingyuan

An erratum was issued for:&#160;<a href="https://jove.unicartagenaproxy.elogim.com/t/64934">Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae</a>. The Authors section was updated.

Erratum: Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

Electron microscopy is one of the most ideal methods for studying the ultrastructure of biological materials that can visually and accurately demonstrate the internal structure of cells at the nanoscale level1. However, due to the complexity of the sample preparation process and the high cost, electron microscopy is not as popular as light microscopy. Recent advancements in electron microscopy techniques have led to significant improvements in image quality, coinciding with a remarkable reduction in the associated workload. Consequently, electron microscopy has assumed an important role in advancing scientific knowledge in diverse fields2.
Drosophila is an excellent animal model for performing genetic manipulations to precisely control the spatial and temporal expression of target genes3. Besides, Drosophila has the advantages of a short growth period and easy rearing compared to mammalian models; therefore, Drosophila is widely used in morphology research4,5.
In Drosophila larvae, neuromuscular junction (NMJ) boutons are widely distributed in the muscles6,7, and immunostaining of NMJ can easily provide information on synapse quantity and morphology8,9 . The NMJ boutons located in the 6th/7th muscles of the A2 and A3 segments are well suited for quantitative and morphological research using light microscopy. This is because of their size and abundance10,11. Therefore, Drosophila larvae NMJs are considered a useful model for neuroscience research12. 
However, it is challenging to observe the NMJ bouton ultrastructure by the TEM. Since the scan window of transmission electron microscopy is narrow, it is hard to position the widely distributed NMJ boutons13. The other reason is that the Drosophila body wall is susceptible to curling during the alcohol dehydration step of the sample preparation protocol7.
Traditional studies have usually chosen boutons between the 6th and 7th muscles of the A2 and A3 segments as sample materials because of their abundance and size14,15. The 6th and 7th muscles of the A2 and A3 segments are bigger than the other muscles and contain more boutons. However, when samples were prepared for electron microscopy, fixed samples tended to become thin and prone to curling, thereby leading to improper positioning of the 6th and 7th muscles of the A2 and A3 segments.
We hereby report a new processing procedure that is more effective in preventing the curling of the samples compared to the traditional method of sample preparation7,16, by allowing the samples to stay flat during the subsequent dehydration, thereby facilitating better positioning of the Drosophila larval neuromuscular junctions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1,2-Epoxypropane</td>
			<td>SHANGHAI LING FENG CHEMICAL REAGENT CO., LTD</td>
			<td>JYJ 037-2015</td>
			<td>Penetrating Agent</td>
		</tr>
		<tr>
			<td>Drosophila Stocks</td>
			<td>Bloomington</td>
			<td>none</td>
			<td>The wild-type control Drosophila strains used in this research were all W1118, and were reared according to standard culture methods</td>
		</tr>
		<tr>
			<td>Flat-bottomed glass test tubes</td>
			<td>Haimen Chenxing Experimental equipment Company&#160;</td>
			<td>none</td>
			<td>Flat-bottomed glass test tubes(bottle)with sponge plug(or bottle stopper)</td>
		</tr>
		<tr>
			<td>K4M cross-linker</td>
			<td>&#160;Agar Scientific</td>
			<td>Cat# 1924B</td>
			<td>The embedding resins are based on a highly cross-linked acrylate and methacrylate formula&#160;</td>
		</tr>
		<tr>
			<td>K4M resin (monomer B)</td>
			<td>&#160;Agar Scientific</td>
			<td>Lot# 631557</td>
			<td>Resin Monomer</td>
		</tr>
		<tr>
			<td>Polyvinyl film</td>
			<td>Haimen Chenxing Experimental equipment Company&#160;</td>
			<td>none</td>
			<td>Transparent polyethylene film is the best , thickness of about 0.2mm</td>
		</tr>
		<tr>
			<td>SPI Chem DDSA</td>
			<td>SPI</td>
			<td>SpI#02827-AF</td>
			<td>Dodecenyl Succinic Anhydride</td>
		</tr>
		<tr>
			<td>SPI-Chem DMP-30 Epoxy</td>
			<td>SPI</td>
			<td>02823-DA</td>
			<td>Accelerator</td>
		</tr>
		<tr>
			<td>SPI-Chem NMA</td>
			<td>SPI</td>
			<td>SpI#02828-AF</td>
			<td>Hardner for Epoxy</td>
		</tr>
		<tr>
			<td>SPI-PON 812 Epoxy</td>
			<td>SPI</td>
			<td>SPI#0259-AB</td>
			<td>Resin Monomer</td>
		</tr>
		<tr>
			<td>Steel mesh&#160;</td>
			<td>Yuhuiyuan Gardening Store(online)</td>
			<td>none</td>
			<td>Copper or stainless steel net</td>
		</tr>
		<tr>
			<td>Transmission electron microscopy</td>
			<td>Hitachi H-7650</td>
			<td>11416692</td>
			<td>All grids (on which samples were gathered) were stained with lead citrate and observed under a transmission electron microscopy.</td>
		</tr>
		<tr>
			<td>Ultrathin microtome</td>
			<td>Leica UC7 ultrathin microtome</td>
			<td>595915</td>
			<td>All sectioning operations are carried out on a Leica UC7 ultrathin microtome using a diamond knife</td>
		</tr>
	</tbody>
</table>

optimizing sample preparation process for transmission electron microscopy of neuromuscular junctions in drosophila larvae

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at the Drosophila NMJ enables researchers to acquire synaptic information, encompassing both quantitative data on synapse abundance and detailed insights into their morphology. However, the diffuse distribution and limited visual range of the TEM present challenges for the ultrastructural analysis. This study introduces an innovative and efficient sample preparation method that surpasses the conventional approach. The procedure begins by placing a metal mesh at the base of a flat-bottomed bottle or test tube, followed by positioning fixed larvae samples onto the mesh. An additional mesh is placed over the samples, ensuring that they are positioned between the two meshes. The fixed samples are thoroughly dehydrated and infiltrated before proceeding with the embedding procedure. Then embedding of the samples in epoxy resin is performed in a flat sheet manner, which allows for the preparation of muscles for positioning and sectioning. Benefiting from these steps, all the muscles of Drosophila larvae can be visualized under light microscopy, therey facilitating subsequent positioning and sectioning. Excess resin is removed after locating the 6th and 7th muscles of body segments A2 and A3. Serial ultra-thin sectioning of the 6th or 7th muscle is performed.

The Drosophila larva body wall is composed of 30 identifiable muscle fibers arranged in a regular pattern and looks like a thin slice after dissection and fixation21(Figure 1A). The sample remains flat during the dehydration process due to the presence of the metal meshes (Figure 1B, C). The larval body muscle is buried in a thin plate made of epoxy resin (Figure 1D-...

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Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

This report provides a new sample preparation procedure for visualizing neuromuscular junctions in Drosophila Larvae. This method is more effective in preventing the curling of the samples compared to the traditional method and is particularly useful for Drosophila neuromuscular junction ultrastructural analysis.

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at ...

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244 Views.  Southeast University. This report provides a new sample preparation procedure for visualizing neuromuscular junctions in Drosophila Larvae. This method is more effective in preventing the curling of the samples compared to the traditional method and is particularly useful for Drosophila neuromuscular junction ultrastructural analysis.

Video: Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron microscope (FIB/SEM). The samples are fixed with aldehydes, heavy metal stained using osmium tetroxide and uranyl acetate. They are then dehydrated with alcohol and infiltrated with resin, which is then hardened. Using a light microscope and ultramicrotome with glass knives, a small block containing the region interest close to the surface is made. The block is then placed inside the FIB/SEM, and the ion beam used to roughly mill a vertical face along one side of the block, close to this region. Using backscattered electrons to image the underlying structures, a smaller face is then milled with a finer ion beam and the surface scrutinised more closely to determine the exact area of the face to be imaged and milled. The parameters of the microscope are then set so that the face is repeatedly milled and imaged so that serial images are collected through a volume of the block. The image stack will typically contain isotropic voxels with dimenions as small a 4 nm in each direction. This image quality in any imaging plane enables the user to analyse cell ultrastructure at any viewing angle within the image stack.

This protocol describes how resin embedded brain tissue can be prepared and imaged in the three dimensions in the focussed ion beam, scanning electron microscope.

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Appetitive Associative Olfactory Learning in Drosophila Larvae

In the following we describe the methodological details of appetitive associative olfactory learning in Drosophila larvae. The setup, in combination with genetic interference, provides a handle to analyze the neuronal and molecular fundamentals of specifically associative learning in a simple larval brain.
Organisms can use past experience to adjust present behavior. Such acquisition of behavioral potential can be defined as learning, and the physical bases of these potentials as memory traces1-4. Neuroscientists try to understand how these processes are organized in terms of molecular and neuronal changes in the brain by using a variety of methods in model organisms ranging from insects to vertebrates5,6. For such endeavors it is helpful to use model systems that are simple and experimentally accessible. The Drosophila larva has turned out to satisfy these demands based on the availability of robust behavioral assays, the existence of a variety of transgenic techniques and the elementary organization of the nervous system comprising only about 10,000 neurons (albeit with some concessions: cognitive limitations, few behavioral options, and richness of experience questionable)7-10.
Drosophila larvae can form associations between odors and appetitive gustatory reinforcement like sugar11-14. In a standard assay, established in the lab of B. Gerber, animals receive a two-odor reciprocal training: A first group of larvae is exposed to an odor A together with a gustatory reinforcer (sugar reward) and is subsequently exposed to an odor B without reinforcement 9. Meanwhile a second group of larvae receives reciprocal training while experiencing odor A without reinforcement and subsequently being exposed to odor B with reinforcement (sugar reward). In the following both groups are tested for their preference between the two odors. Relatively higher preferences for the rewarded odor reflect associative learning - presented as a performance index (PI). The conclusion regarding the associative nature of the performance index is compelling, because apart from the contingency between odors and tastants, other parameters, such as odor and reward exposure, passage of time and handling do not differ between the two groups9.

Appetitive Associative Olfactory Learning in Drosophila Larvae

Drosophila larvae are able to associate odor stimuli with gustatory reward. Here we describe a simple behavioral paradigm that allows the analysis of appetitive associative olfactory learning.

In the following we describe the methodological  details of appetitive associative olfactory learning in Drosophila larvae. The setup, in combination with ...

Foraging Path-length Protocol for Drosophila melanogaster Larvae

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral variation. The larval path-length assay was developed to measure individual differences in foraging behavior that were later linked to the foraging gene. Larval path-length is an easily scored trait that facilitates the collection of large sample sizes, at minimal cost, for genetic screens. Here we provide a detailed description of the current protocol for the larval path-length assay first used by Sokolowski. The protocol details how to reproducibly handle test animals, perform the behavioral assay and analyze the data. An example of how the assay can be used to measure behavioral plasticity in response to environmental change, by manipulating feeding environment prior to performing the assay, is also provided. Finally, appropriate test design as well as environmental factors that can modify larval path-length such as food quality, developmental age and day effects are discussed.

Foraging Path-length Protocol for Drosophila melanogaster Larvae

We provide a detailed protocol for a Drosophila melanogaster foraging path-length assay. We discuss the preparation and handling of test animals, how to perform the assay and analyze the data.

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral ...

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and characterize ultrastructural and morphological details of neurons, such as synaptic contacts. Good preservation of brain tissue for electron microscopy can be obtained by rigorous cryo-fixation methods, but these techniques are rather costly and limit the use of immunolabeling, which is crucial to understand the connectivity of identified neuronal systems. Freeze-substitution methods have been developed to allow the combination of cryo-fixation with immunolabeling. However, the reproducibility of these methodological approaches usually relies on costly freezing devices. Moreover, achieving reliable results with this technique is very time-consuming and skill-challenging. Hence, the traditional chemically fixed brain, particularly with acrolein fixative, remains a time-efficient and low-cost method to combine electron microscopy with immunohistochemistry. Here, we provide a reliable experimental protocol using chemical acrolein fixation that leads to the preservation of primate brain tissue and is compatible with pre-embedding immunohistochemistry and transmission electron microscopic examination.

Here, we provide an easy, low-cost, and time-efficient protocol to chemically fix primate brain tissue with acrolein fixative, allowing for long-term preservation that is compatible with pre-embedding immunohistochemistry for transmission electron microscopy.

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and ...

Novel Assay for Cold Nociception in Drosophila Larvae

How organisms sense and respond to noxious temperatures is still poorly understood. Further, the mechanisms underlying sensitization of the sensory machinery, such as in patients experiencing peripheral neuropathy or injury-induced sensitization, are not well characterized. The genetically tractable Drosophila model has been used to study the cells and genes required for noxious heat detection, which has yielded multiple conserved genes of interest. Little is known however about the cells and receptors important for noxious cold sensing. Although, Drosophila does not survive prolonged exposure to cold temperatures (&#8804;10 &#186;C), and will avoid cool, preferring warmer temperatures in behavioral preference assays, how they sense and possibly avoid noxious cold stimuli has only recently been investigated.
Here we describe and characterize the first noxious cold (&#8804;10 &#186;C) behavioral assay in Drosophila. Using this tool and assay, we show an investigator how to qualitatively and quantitatively assess cold nociceptive behaviors. This can be done under normal/healthy culture conditions, or presumably in the context of disease, injury or sensitization. Further, this assay can be applied to larvae selected for desired genotypes, which might impact thermosensation, pain, or nociceptive sensitization. Given that pain is a highly conserved process, using this assay to further study&#160;thermal nociception will likely glean important understanding of pain processes in other species, including vertebrates.

Novel Assay for Cold Nociception in Drosophila Larvae

Here we demonstrate a novel assay to study cold nociception in Drosophila larvae. This assay utilizes a custom-built Peltier probe capable of applying a focal noxious cold stimulus and results in quantifiable cold-specific behaviors. This technique will allow further cellular and molecular dissection of cold nociception.

How organisms sense and respond to noxious temperatures is still poorly understood. Further, the mechanisms underlying sensitization of the sensory ...

Sample Preparation for Endopeptidomic Analysis in Human Cerebrospinal Fluid

This protocol describes a method developed to identify endogenous peptides in human cerebrospinal fluid (CSF). For this purpose, a previously developed method based on molecular weight cut-off (MWCO) filtration and mass spectrometric analysis was combined with an offline high-pH reverse phase HPLC pre-fractionation step.
Secretion into CSF is the main pathway for removal of molecules shed by cells of the central nervous system. Thus, many processes in the central nervous system are reflected in the CSF, rendering it a valuable diagnostic fluid. CSF has a complex composition, containing proteins that span a concentration range of 8 - 9 orders of magnitude. Besides proteins, previous studies have also demonstrated the presence of a large number of endogenous peptides. While less extensively studied than proteins, these may also hold potential interest as biomarkers.
Endogenous peptides were separated from the CSF protein content through MWCO filtration. By removing a majority of the protein content from the sample, it is possible to increase the sample volume studied and thereby also the total amount of the endogenous peptides. The complexity of the filtrated peptide mixture was addressed by including a reverse phase (RP) HPLC pre-fractionation step at alkaline pH prior to LC-MS analysis. The fractionation was combined with a simple concatenation scheme where 60 fractions were pooled into 12, analysis time consumption could thereby be reduced while still largely avoiding co-elution.
Automated peptide identification was performed by using three different peptide/protein identification software programs and subsequently combining the results. The different programs were complementary rather than comparable with less than 15% of the identifications overlapped between the three.

A method for mass spectrometric analysis of endogenous peptides in human cerebrospinal fluid (CSF) is presented. By employing molecular weight cut-off filtration, chromatographic pre-fractionation, mass spectrometric analysis and a subsequent combination of peptide identification strategies, it was possible to expand the known CSF peptidome nearly ten-fold compared to previous studies.

This protocol describes a method developed to identify endogenous peptides in human cerebrospinal fluid (CSF). For this purpose, a previously developed ...

Levator Auris Longus Preparation for Examination of Mammalian Neuromuscular Transmission Under Voltage Clamp Conditions

This protocol describes a technique to record synaptic transmission from the neuromuscular junction under current-clamp and voltage-clamp conditions. An ex vivo preparation of the levator auris longus (LAL) is used because it is a thin muscle that provides easy visualization of the neuromuscular junction for microelectrode impalement at the motor endplate. This method allows for the recording of spontaneous miniature endplate potentials and currents (mEPPs and mEPCs), nerve-evoked endplate potentials and currents (EPPs and EPCs), as well as the membrane properties of the motor endplate. Results obtained from this method include the quantal content (QC), number of vesicle release sites (n), probability of vesicle release (prel), synaptic facilitation and depression, as well as the muscle membrane time constant (&#964;m) and input resistance. Application of this technique to mouse models of human disease can highlight key pathologies in disease states and help identify novel treatment strategies. By fully voltage-clamping a single synapse, this method provides one of the most detailed analyses of synaptic transmission currently available.

Levator Auris Longus Preparation for Examination of Mammalian Neuromuscular Transmission Under Voltage Clamp Conditions

The protocol described in this paper uses the mouse levator auris longus (LAL) muscle to record spontaneous and nerve-evoked postsynaptic potentials (current-clamp) and currents (voltage-clamp) at the neuromuscular junction. Use of this technique can provide key insights into mechanisms of synaptic transmission under normal and disease conditions.

This protocol describes a technique to record synaptic transmission from the neuromuscular junction under current-clamp and voltage-clamp conditions. An ...

Preparation of the Rat Vocal Fold for Neuromuscular Analyses

The purpose of this tutorial is to describe the preparation of the rat vocal fold for histochemical neuromuscular study. This protocol outlines procedures for rat laryngeal dissection, flash-freezing, and cryosectioning of the vocal folds. This study describes how to cryosection vocal folds in both longitudinal and cross-sectional planes. A novelty of this protocol is the laryngeal tracking during cryosectioning that ensures accurate identification of the intrinsic laryngeal muscles and reduces the chance of tissue loss. Figures demonstrate the progressive cryosectioning in both planes. Twenty-nine rat hemi-larynges were cryosectioned and tracked from the emergence of the thyroid cartilage to the appearance of the first section that included the full vocal fold. The full vocal fold was visualized for all animals in both planes. There was high variability in the distance from the appearance of the thyroid cartilage to the appearance of the full vocal fold in both planes. Weight was not correlated to depth of laryngeal landmarks, suggesting individual variability and other factors related to tissue preparation may be responsible for the high variability in the appearance of landmarks during sectioning. This study details a methodology and presents morphological data for preparing the rat vocal fold for histochemical neuromuscular investigation. Due to high individual variability, laryngeal landmarks should be closely tracked during cryosectioning to prevent oversectioning tissue and tissue loss. The use of a consistent methodology, including adequate tissue preparation and awareness of landmarks within the rat larynx, will assist with consistent results across studies and aid new researchers interested in using the rat vocal fold as a model to investigate laryngeal neuromuscular mechanisms.

This protocol describes methods used to prepare rat vocal folds for histochemical neuromuscular study.

The purpose of this tutorial is to describe the preparation of the rat vocal fold for histochemical neuromuscular study. This protocol outlines procedures ...

Dissection of Single Skeletal Muscle Fibers for Immunofluorescent and Morphometric Analyses of Whole-Mount Neuromuscular Junctions

The neuromuscular junction (NMJ) is a specialized point of contact between the motor nerve and the skeletal muscle. This peripheral synapse exhibits high morphological and functional plasticity. In numerous nervous system disorders, NMJ is an early pathological target resulting in neurotransmission failure, weakness, atrophy, and even in muscle fiber death. Due to its relevance, the possibility to quantitatively assess certain aspects of the relationship between NMJ components can help to understand the processes associated with its assembly/disassembly. The first obstacle when working with muscles is to gain the technical expertise to quickly identify and dissect without damaging their fibers. The second challenge is to utilize high-quality detection methods to obtain NMJ images that can be used to perform quantitative analysis. This article presents a step-by-step protocol for dissecting extensor digitorum longus and soleus muscles from rats. It also explains the use of immunofluorescence to visualize pre and postsynaptic elements of whole-mount NMJs. Results obtained demonstrate that this technique can be used to establish the microscopic anatomy of the synapsis and identify subtle changes in the status of some of its components under physiological or pathological conditions.

The ability to accurately detect neuromuscular junction components is crucial in evaluating modifications in its architecture because of pathological or developmental processes. Here we present a complete description of a straightforward method to obtain high-quality images of whole-mount neuromuscular junctions that can be used to perform quantitative measurements.

The neuromuscular junction (NMJ) is a specialized point of contact between the motor nerve and the skeletal muscle. This peripheral synapse exhibits high ...

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

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Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Appetitive Associative Olfactory Learning in Drosophila Larvae

Foraging Path-length Protocol for Drosophila melanogaster Larvae

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Novel Assay for Cold Nociception in Drosophila Larvae

Sample Preparation for Endopeptidomic Analysis in Human Cerebrospinal Fluid

Levator Auris Longus Preparation for Examination of Mammalian Neuromuscular Transmission Under Voltage Clamp Conditions

Preparation of the Rat Vocal Fold for Neuromuscular Analyses

Dissection of Single Skeletal Muscle Fibers for Immunofluorescent and Morphometric Analyses of Whole-Mount Neuromuscular Junctions