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

Summary

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.

Abstract

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 6^th and 7^th muscles of body segments A₂ and A₃. Serial ultra-thin sectioning of the 6^th or 7^th muscle is performed.

Introduction

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 research^4,5.

In Drosophila larvae, neuromuscular junction (NMJ) boutons are widely distributed in the muscles^6,7, and immunostaining of NMJ can easily provide information on synapse quantity and morphology^8,9 . The NMJ boutons located in the 6^th/7^th 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 abundance^10,11. 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 6^th and 7^th muscles of the A₂ and A₃ segments as sample materials because of their abundance and size^14,15. The 6^th and 7^th 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 6^th and 7^th 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 preparation^7,16, by allowing the samples to stay flat during the subsequent dehydration, thereby facilitating better positioning of the Drosophila larval neuromuscular junctions.

Protocol

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

1. Dissection
   1. Dissect the wandering late-3rd-instar larvae in Jan solution and obtain the entire body wall based on the standard procedures¹⁷. Prepare Jan'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.
2. Fixation
   1. Fix the larval body wall in the dissection chamber at 4 °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)
   2. Rinse the samples several times with sodium cacodylate buffer.
   3. 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.
   4. Stain the Drosophila neuromuscular junction samples with 2% saturated uranyl acetate for 2 h, followed by several rinses with deionized water (Figure 1A).
3. Sample placement
   1. Cut two round pieces of metal mesh of similar size (pore size 270 µm) with scissors.
   2. Place a metal mesh (270 µm) at the base of a flat-bottomed bottle in advance.
   3. 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.

2. Dehydration

NOTE In this paper, the ethanol gradient dehydration method was used.

1. 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 °C.
2. Rinse the samples twice with propylene oxide at room temperature for 5 min each time (Figure 1C).

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.

1. 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.

4. Embedding

1. Prepare a polyethylene film and cut it into a big square (approximately 5 × 5 cm) and a small hollow rectangular support (approximately 2 × 2 cm). Glue the small square with the big square.
2. Pre-position sufficient epoxy resin at the center of the big square.
3. Place the samples in the pre-positioned epoxy resin with the muscle side facing upwards. Use a stereoscope.
4. Add several drops of resin to the samples and then cover them with PE film (a little larger than the spacer) (Figure 1D-E).
5. Polymerize the samples at 37 °C, 42 °C and 60 °C for 24 h respectively (Figure 1F).

5. Sectioning procedure

1. 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 13^th 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.
2. Remove the trapezoid from the remainder of the sample and adhere it to the resin-filled capsule with the A/B glue.
3. Confirm the position of the 6^th and 7^th muscles of A₂ and A₃ segments under the light microscopy (Figure 1G) and maintain the tangent plane parallel to the 6^th muscle.
4. Cut away one-fifth of the sample along the two sides of the A₃ segment.
   NOTE: Remove most of the 6^th muscle, and retain 1/3 of the width of the 7^th muscle.
5. Using an ultrathin microtome, prepare an ultrathin section of the sample starting with the 6^th muscle.
   NOTE: Each slice is approximately 90 nm thick.
6. Attached as many slices as possible to each copper mesh.
7. Perform transmission electron microscopy as described previously¹⁶.

Results

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-...

Discussion

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...

Materials

List of materials used in this article

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