Validation of Hyperbaric Pressure System with Xenon Anesthesia for Drosophila melanogaster

Summary

Gases that seem harmless at atmospheric pressure can induce behaviors like narcosis under hyperbaric conditions. Conventional hyperbaric pressure chambers are costly and labor-intensive. This study presents a straightforward, low-cost method for examining xenon's effects on Drosophila melanogaster at moderate pressures below 4 atm.

Abstract

Under hyperbaric conditions, the elevated concentrations of the ambient gases may elicit a range of behavioral responses in various animals. A substantial portion of these behaviors represents changes in cognitive states that are yet poorly understood. One notable example is general anesthesia. The behavioral effect of several gases under hyperbaric conditions in general anesthesia, or narcosis, has been a long-lasting scientific inquiry, despite the overwhelmingly frequent use of the drugs in medicine. Xenon, classified as a noble gas and an anesthetic agent, is regarded as one of the safest options for general anesthesia in humans. However, it does not anesthetize flies under normobaric conditions. Here, we established a simple experimental setup that increases the ambient pressure up to ~3 atm and allows the study of xenon anesthesia in one of the most popular model organisms, Drosophila melanogaster. This experimental setup further facilitates the study of other gases and their effect under hyperbaric conditions.

Introduction

The two extreme ends of behavioral effects that have been observed in animals under hyperbaric conditions are convulsion by oxygen and narcosis by nitrogen^1,2. Curiously, cognitive impairments such as hallucinations and memory impairment have been reported with nitrogen and hydrogen at pressure values prior to the level of unconsciousness^3,4. Anesthesia induced by xenon, krypton, and nitrous oxide under hyperbaric conditions has been reported before^5,6.

General anesthesia is both a cornerstone of medicine and a phenomenon of considerable scientific interest. Despite its long and widespread use, our understanding remains limited about how exactly some chemicals induce the temporary absence of movement, memory, pain, and consciousness⁷. Xenon, being a noble gas, could be appointed as the most peculiar member among anesthetic agents. It is regarded as a safe option for anesthesia, offering rapid induction and recovery, minimal physiological disturbance, and only mild side effects such as postoperative nausea^8,9.

Given the medical importance of anesthesia and the unusual chemical properties of xenon, this study aims to investigate xenon-induced anesthesia using the simple animal model Drosophila melanogaster. A variety of research studies have been conducted using fruit flies to study general anesthesia, with many proposed setups and the use of various volatile anesthetics^10,11,12,13. However, there has been no report of xenon anesthesia in Drosophila. Under normobaric conditions, xenon does not induce anesthesia in Drosophila melanogaster, commonly known as fruit flies. To address this limitation, this study offers a hyperbaric system that allows controlled manipulation of ambient pressure between 1 atm and approximately 4 atm. There have been several studies using hyperbaric chambers to answer similar curiosities^14,15. However, most commercial hyperbaric chambers benefit from sturdy, leak-proof materials, but they have a high volume (~3 L) and are systematically complicated to operate and expensive. Instead, the method described in this article is simple and inexpensive, allowing for the exploration of the effects of various gases and their combinations. It provides a setup that is suitable for small animal models like D. melanogaster and C. elegans.

Protocol

1. Drosophila melanogaster preparation

1. Collect 20 female and 20 male flies aged 1-3 days for each experiment using CO₂.
2. Rest the selected flies in a vial with food for at least 24 h prior to the experiment at a temperature of ~22 °C.

2. Anesthetic delivery

1. Transfer the gases from their container gas cylinders to the leakproof gas bags for experimental use. For the direct transfer of gas from the container to the bags, use silicone tubing.
2. Attach 25 mm (3.4 mm diameter) silicone tubing to the polypropylene syringe. Then connect the polypropylene syringe to the gas bags and to the three-way tap. For clarity, the polypropylene syringe will be referred to as syringe #1 in the remainder of the text.

3. Assembly of the hyperbaric setup (Figure 1)

1. Position the LED light table in landscape orientation on the table (long edge horizontal to the table surface).
2. Place the injection pump adjacent to one of the short sides of the LED light table, ensuring that the pump's orientation permits the experimental syringe to extend above the backlight.
3. Locate the camera above the setup to cover the region that the experimental syringe resides in.
4. Cut a 75 mm long silicon tube (3.4 mm diameter). Connect the three-way tap to the manometer using the silicone tubing from the central nozzle.
5. Use hose spring clamps on the nozzle from the side of the manometer and the three-way tap to ensure that the tubing is tight.
6. Using the button marked by a double arrow in the control unit of the injection pump, set the speed to 80 mm/m and duration to 38 s for the automatic compression or decompression of the plunger.

4. Procedure for Xenon anesthesia

1. Limit the ambient light in the experiment room and turn on the LED light table and set the brightness at level 3. This helps increase the contrast in the video.
2. Remove the plunger of the polycarbonate syringe. Insert a small amount of cotton in the nipple of the syringe to stop flies from escaping. For clarity, the polycarbonate syringe will be referred to as syringe #2 in the remainder of the text.
3. To transfer 40 flies from their vial to syringe #2, first take out the lid of the vial and place the syringe barrel vertically on top of the vial. The barrel flange should merge with the top of the vial to allow the flies to enter the syringe barrel. Turn the construct 180° and tap. This will place the flies at the bottom of the syringe barrel. Place the plunger back in. Place the syringe with the flies inside the injection pump.
4. Orient the barrel marks of the syringe on the side for a better view of the flies from the camera's point of view.
5. Secure the syringe from the barrel and the plunger head to the injection pump using the adjustable screws.
6. Remove the cotton from the tip of syringe #2 and connect it with the 3-way tap.
7. Start the video recording. Turn the manometer on. Turn the stopcock to connect the manometer with syringe #2 and confirm 0 kPa pressure in the syringe using the manometer.
8. Turn the 3-way tap to the lateral position to allow air exchange with the syringe prior to the experiment.
9. Move the plunger from the 20 to the 2 mL mark via the manual control option of the injection pump, which is designed to facilitate the quickest option.
10. Aspirate 18 mL of the anesthetic gas from the gas bag to syringe #1. Connect syringe #1 with the three-way tap from the lateral luer lock port. To assess the correct volume, 18 mL of anesthetic gas in a total amount of 20 mL mixture would give 90% of the gas percentage. This can be changed according to the desired final gas percentage.
11. Move the plunger back to the 20 mL line by the manual option of the injection pump.
12. Turn the stopcock to connect the manometer with syringe #2. Usually, the manometer shows ~-3 kPa pressure after the filling of the anesthetic.
13. Move the plunger automatically, with speed and time settings set to 80 mm/s and 38 s. The pump is expected to reach the 5 mL bar of the barrel in 38 s. When the plunger stops, the maximum pressure must be ~200 kPa.
14. Leave the plunger at the 5 mL position for 20 s. Note the peak pressure.
15. Tap the syringe to dislodge any flies clinging to the plunger, making them settle motionless at the bottom.
16. Start the automatic decompression at the end of 20 s. Terminate the experiment when the plunger is back at the 20 mL bar. Note the final pressure for the leak control.

5. Data analysis

NOTE: In this experiment, anesthetic emergence was used as an endpoint. The fly movements were quantified from the start time of plunger decompression. Two different analysis methods were developed: pixel analysis and fly tracking. Each analysis method had its drawbacks, which are communicated in the discussion.

1. Pixel analysis
   1. Use ScreenFlow to edit the videos. Crop each video to the region of interest (ROI), i.e., first 5 mL of the syringe barrel.
   2. Trim the video to the first 20 s after syringe decompression. Export the edited videos as GIFs. Duplicate the GIF.
   3. Using Fiji, remove the first frame from one duplicate and remove the last frame from the other duplicate. To achieve this, select Image > Video Editing > Delete Frames.
   4. Make each GIF binary in Fiji so flies appear as white on a black background. To achieve this, select Process > Binary > Make binary.
   5. Open both GIFs in Fiji. Calculate the pixel difference between the GIFs. To achieve this, select Process > Image Calculator. This operation will create a pixel difference GIF.
   6. Save the pixel difference GIF. Analyze the particle in the different GIFs. To do this, select Analyze > Analyze particles in Fiji. Adjust pixel size thresholds to increase the signal-to-noise ratio.
   7. After analyzing particles, Fiji outputs a CSV with pixel differences per frame. Save the CSV. CSV data can be used to plot the movement per frame. Use cumulative movement through frames to assign statistical differences between the experiments.
2. Fly tracking
   NOTE: For the fly tracking method, AI technology was utilized to detect and track the center of mass of each fly in the experiment videos. Two models were combined for this: YOLO (You Only Look Once) for detection and MASA (Matching Anything by Segmenting Anything) for tracking. The models were executed on Google Colab using a Tesla T4 GPU.
   1. Install the Ultralytics YOLO library in Colab, then load the yolo11n.pt model. Use CVAT (Computer Vision Annotation Tool) to annotate flies in frames extracted from the videos, then export the dataset in YOLO format for training. Detailed installation and training procedures are described in the online repository.
   2. Install the MASA repository in Google Colab and incorporate a Kalman filter to perform object tracking across consecutive frames. Integrate the trained YOLO detection model with the MASA tracking framework.
   3. Use FixTrack to manually review the tracklets generated by the combined MASA+YOLO pipeline. Identify and merge incorrectly fragmented tracklets or correct identity mismatches to obtain continuous trajectories for individual flies. Detailed installation and training procedures are described in the FixTrack repository.

Results

The method described in this paper allows for the monitoring of both the timing and extent of locomotor activity in flies under hyperbaric conditions. This configuration permits gases like xenon and nitrous oxide, which do not have anesthetic properties under normal conditions, to produce anesthesia in Drosophila melanogaster. To demonstrate this, we experimented with different concentrations of xenon and air. We began establishing the process with 90% xenon and 10% air. Then we demonstrated the methodology's se...

Discussion

This simple, low-cost setup makes hyperbaric behavioral assays in Drosophila broadly accessible, enabling rapid, reproducible experiments without specialized pressure vessels. However, it presents a few limitations. One of its important compromises is the lower maximum pressure relative to existing setups. The highest pressure we can induce with this setup is ~4 atm, whereas pressure vessels used for Drosophila are reported to reach roughly double that, and systems for mice can go up to 100 atm

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Airtight gas bags (0.5 L)	Techinstro	7440440
Camera	Apple	Iphone 15
Double wire spring hose clamps	sourcing map	a24120200ux2109
Drosophila food mix	Advanced Husbandary	DM900
Drosophila melanogaster	Darwin Biological	LZ-210
Injection pipe	Aitoserlea	B09M8G5BQ8
LED light table	Lairi	M1-3878368
Manometer	Dwyeromega	477AV-8
Metal three-way compression nut stopcock	Sigma Aldrich	Z182184
Polycarbonate syringe (20 mL)	DMC Medical	FD-1031NS
polypropylene syringe (20 mL)	BD	301031
Silicone tubing
Xenon Gas	Chengdu Taiyu Industrial Gases