Controlled Semi-Automated Laser-Induced Injuries for Studying Spinal Cord Regeneration in Zebrafish Larvae

Summary

The present protocol describes a method to induce tissue-specific and highly reproducible injuries in zebrafish larvae using a laser lesion system combined with an automated microfluidic platform for larvae handling.

Abstract

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes this animal model very useful for studying spinal cord injury and regeneration. The standard protocol for inducing such lesions is to transect the dorsal part of the trunk manually. However, this technique requires extensive training and damages additional tissues. A protocol was developed for laser-induced lesions to circumvent these limitations, allowing for high reproducibility and completeness of spinal cord transection over many animals and between different sessions, even for an untrained operator. Furthermore, tissue damage is mainly limited to the spinal cord itself, reducing confounding effects from injuring different tissues, e.g., skin, muscle, and CNS. Moreover, hemi-lesions of the spinal cord are possible. Improved preservation of tissue integrity after laser injury facilitates further dissections needed for additional analyses, such as electrophysiology. Hence, this method offers precise control of the injury extent that is unachievable manually. This allows for new experimental paradigms in this powerful model in the future.

Introduction

In contrast to mammals, zebrafish (Danio rerio) can repair their central nervous system (CNS) after injury¹. The use of zebrafish larvae as a model for spinal cord regeneration is relatively recent. It has proven valuable to investigate the cellular and molecular mechanisms underlying repair². This is due to the ease of manipulation, the short experimental cycle (new larvae every week), the tissues' optical transparency, and the larvae's small size, ideally suited for in vivo fluorescence microscopy.

In the case of spinal cord regeneration, two additional advantages of using larvae are the speed of recovery, a few days compared to a few weeks for adults, and the ease of inducing injuries using manual techniques. This has been successfully used in many studies^3,4,5, including recent investigations^6,7. Overall, this leads to increased meaningful data production, high adaptability of experimental protocols, and decreased experimental costs. The use of larvae younger than 5 days post-fertilization also reduces the use of animals following the 3R principles in animal research⁸.

After a spinal cord injury in zebrafish larvae, many biological processes occur, including inflammatory response, cell proliferation, neurogenesis, migration of surviving or newly generated cells, reformation of functional axons, and a global remodeling of neural processes circuits and spine tissues^6,7,9,10. To be successfully orchestrated, these processes involve a finely regulated interaction between a range of cell types, extracellular matrix components, and biochemical signals^11,12. Unraveling the details of this significant reorganization of a complex tissue such as the spinal cord requires the use and development of precise and controlled experimental approaches.

The primary experimental paradigm used to study spinal cord regeneration in zebrafish is to use surgical means to induce tissue damage by resection, stabbing, or cryoinjury^3,13. These approaches have the disadvantage of requiring specific training in microsurgery skills, which is time-consuming for any new operator and may prevent their use in short-term projects. Furthermore, they usually induce damage to the surrounding tissues, which may influence regeneration.

Another approach is to induce cell damage chemically¹⁴ or by genetic manipulations¹⁵. The latter allows for highly targeted damage. However, such a technique requires long preparatory work to generate new transgenic fish before doing any experiment, renewed each time a unique cell type is targeted.

There is, thus, the need for a method allowing targeted but versatile lesions suitable to a variety of studies in regeneration. A solution is to use a laser to induce localized damage in the tissue of interest^{16,17,18,19,20}. Indeed, the use of laser-induced tissue damage presents a robust approach for generating spinal cord lesions with many advantages. The microscopes equipped with such laser manipulation modules allow specifying a customized shaped area where cell ablation will occur, with the extra benefit of temporal control. The size and position of the lesion can be thus adapted to address any questions.

The missing feature of most laser lesion systems is the possibility to induce injuries in a highly reproducible way for a series of larvae. Here an original protocol is described using a UV laser to induce semi-automated precise and controlled lesions in zebrafish larvae based on a microfluidic platform designed for automated larvae handling²¹. Moreover, in the system presented here, larvae are inserted in a glass capillary, which permits free rotation of the animal around its rostrocaudal axis. The user can choose which side of the larva to present to the laser while allowing fluorescence imaging to precisely target the laser beam and assess the damage after the lesion.

The protocol described here is used with a semi-automated zebrafish larvae imaging system combined with a spinning disk equipped with a UV laser (designated hereafter as the VAST system). However, the main points of the protocol and most of the claims of the technique are valid for any system equipped with a laser capable of cell ablation, including two-photon laser scanning microscopes, spinning-disk microscopes provided with a UV laser (FRAP module), or video-microscopes with a laser module for photo manipulation. One of the main differences between the VAST system and conventional sample handling will be that for the latter, mounting larvae in low-melting-point agarose on glass coverslips/glass-bottom Petri dishes in place of loading them in a 96-well plate will be required.

The benefits offered by this method open opportunities for innovative research on the cellular and molecular mechanisms during the regeneration process. Moreover, the high data quality allows for quantitative investigations in a multidisciplinary context.

Protocol

All animal studies were carried out with approval from the UK Home Office and according to its regulations, under project license PP8160052. The project was approved by the University of Edinburgh Institutional Animal Care and Use Committee. For experimental analyses, zebrafish larvae up to 5-day-old of either sex were used of the following available transgenic lines: Tg(Xla.Tubb:DsRed;mpeg1:GFP), Tg(Xla.Tubb:DsRed), Tg(betaactin:utrophin-mCherry), Tg(Xla.Tubb:GCaMP6s), and Tg(mnx1:gfp) (see Supplementary File 1 regarding the generation of the transgenic zebrafish lines). A schematic of the protocol using the automated zebrafish larvae handling platform is shown in Figure 1. All custom software, scripts, and detailed experimental protocols used in this work are available at https://github.com/jasonjearly/micropointpy/.

1. Sample preparation

1. At 5 h post-fertilization, sort the embryos for the correct developmental stage²¹. Discard dead eggs and poorly developed and overdeveloped embryos.
2. At 3 days post-fertilization (dpf), anesthetize larvae by adding 2 mL of 0.4% aminobenzoic-acid-ethyl methyl-ester to 50 mL of fish facility water in a 90 mm Petri dish. Use animals raised with phenylthiourea (PTU) (see Table of Material) to prevent skin pigmentation if it is an issue, which is not the case for spinal cord injuries on 3 dpf larvae described in this protocol.
   NOTE: This relatively high anesthetic concentration is used to prevent movements of the larvae following the laser impact.
3. Screen the embryos for fluorescent reporter expression (Supplementary File 1).
   NOTE: A fluorescent reporter for the spinal cord (or other structure of interest) is often required to assess the efficiency of the injury. The use of tg(Xla.Tubb:DsRed) helps to identify the spinal cord.
4. Transfer the selected larvae into a 96-well plate for use in the VAST system (see Table of Materials) with 300 µL of fish facility water per well. Use the medium containing the anesthetic from the 90 mm Petri dish directly. Ensure to have only one larva per well. Prepare one extra empty 96-well plate to collect the lesioned larvae.
   ​NOTE: If using another laser lesion system, mount the larvae in 1% Low-Melting Point (LMP) agarose gel in an appropriate observation chamber.

2. Microscope preparation

1. Switch on all the system components (VAST, microscope, laser, PC), including the laser for ablation.
2. Once the hardware is fully initialized, launch the microscope software, ImageJ/Fiji, a python integrated development environment (IDE), and the automated zebrafish imaging (VAST system) software if using this platform (see Table of Materials).
3. Set up the VAST software following the steps below.
   1. When the VAST software launches, choose Plate on the first window and click on the Done button (Figure 2A). Another small window will pop up asking whether the capillary is empty and clean. Verify by looking at the image of the capillary if there are any air bubbles inside. If not, click on Yes. If there are any bubbles, click on No and follow step 2.3.2-2.3.3 (Figure 2B).
   2. On the Large Particle (LP) Sampler window, click on Prime to remove air bubbles (Figure 2C).
   3. Go to the main software window (with the capillary image) and right-click on the image. Select Record empty capillary image on the pop-up menu (Figure 2B).
   4. In the LP Sampler window, go to the File menu and select the Open Script option. Choose a file containing the script corresponding to the experiment to be performed.
   5. In the main VAST software window, go to File and choose Open Experiment. Choose the experiment file corresponding to the planned experiment.
      NOTE: Ensure that the boxes Auto unload and Bulk output to waste are NOT checked.
4. Set up the microscope software for imaging.
   1. Launch the microscope imaging software (see Table of Materials) to initialize the hardware. This may take a few minutes, depending on the system.
   2. Go to the acquisition settings and set up the microscope for imaging the fluorophore expressed in the larvae. Use a 10x NA 0.5 water-dipping objective to ensure the focal volume is elongated enough along the optical axis to lesion the whole depth of the spinal cord or the targeted tissue.
5. Set up ImageJ/Fiji for laser lesions.
   1. Go to the File menu, choose New/Script to open the script window.
   2. In the New window, go to the File menu and choose Open to load the laser lesion script (Manual_MP_Operation.ijm).
6. Set up the Python IDE.
   1. Launch the Python IDE.
   2. Go to the File menu and choose Open File to load the script to manage the laser (Watch_for_ROIs_py3.py).
   3. Go to the Run menu and choose Run Without Debugging to run the script. Check to ensure that a sequence of messages in the TERMINAL panel appears along with some noise while the laser attenuator initializes (Figure 2D).

3. Performing laser lesions on the VAST system

1. Center the capillary relative to the microscope objective by moving the stage by clicking on the arrow buttons on the main window of the VAST software (Figure 2B).
2. Focus on the top of the capillary by looking through the eyepieces and using the transmitted light of the microscope.
   CAUTION: The capillary is very fragile and may break if touched by the objective. Move the microscope knob slowly when focusing in and out.
3. Place the 96-well plates on the plate holder of the LP Sampler of the VAST system. Place the plate containing larvae on the left holder and the plate for collection on the right. Ensure that the plates are correctly oriented: the A1 well must be in the front-left corner of the holder.
4. In the VAST software, on the LP Sampler window, click on the Plate Template button and select all the wells containing larvae. Click on the OK button to validate and close the window (Figure 2C).
5. In LP Sampler window, click on the Run Plate button to start loading a larva.
   NOTE: After some time, the larva should be visible in the capillary at position (predefined in the experiment definition file), allowing to injure the spinal cord. The VAST tray light will turn off after a few rotations to set the larva with the lateral side facing the microscope objective.
6. Go to the microscope software and click on the Live button to image the larva.
7. Turn the microscope focus knob until the spinal cord central canal is visible.
   NOTE: It can be easier to focus using transmitted light first, and then refine with fluorescence.
8. Take a snapshot in fluorescence and save the image to a dedicated folder.
9. Open the image in ImageJ and adjust the contrast if required (using the Image/Adjust/Brightness/contrast... menu in ImageJ).
10. Click on the Region of Interest (ROI) line tool and draw a short line (20 µm) centered on the spinal cord (Figure 3A).
11. Switch the microscope to the 100% reflective mirror position.
12. Load the ImageJ script and click on the Run button. Use the following parameters: Repetition - 2; Sample - 1; Width - 40-micron; Attenuation - 89 (Full laser power) (Figure 3C).
13. When the laser shot sequence is finished, switch to fluorescence imaging on the imaging software and adjust the focus if required.
    NOTE: A shift in focus is often observed due to tail displacement during laser exposure.
14. Take a new snapshot and save it.
15. Open this new image in ImageJ and draw a new line that should be larger than the spinal cord itself (~80 µm), starting below the ventral side of the spinal cord in the upper part of the notochord and going towards the dorsal side to end in the space between the spinal cord and the skin (Figure 3B).
16. Switch the microscope to the 100% reflective mirror position.
17. Go to the ImageJ script window and click on the Run button. Use the following parameters: Repetition - 2; Sample - 1; Width - 40 microns; Attenuation - 89 (Full laser power).
18. After the (longer) laser shot sequence is finished, verify the transection quality by imaging fluorescence and focusing. Ensure that no cell or axons remain intact in the lesion site, which should appear as a dark or as a faint and homogeneous fluorescent area (Figure 3D, bottom panel).
19. Collect the lesioned larvae into the empty 96-well plate (with the same well co-ordinates as that of the original well) by going to the main VAST software window and clicking on the Collect button.
20. Switch back on the VAST system light by clicking on the check box Tray Light on the bottom left of the window.
21. Repeat step 3.3-3.17 for each new larva to be injured.

4. Post-lesion handling and additional experiments

1. Take out larvae from the 96-well plate as soon as possible and transfer them to a clean Petri dish with fresh fish facility water for the larvae to recover post-lesion. Put the Petri dish in an incubator at 28 °C.
   ​NOTE: The damage often continues to propagate in the first hour after the lesion. The actual extent of the lesion should thus be assessed by fluorescence imaging after a delay of approximately 1 h.

5. Troubleshooting

1. If air bubbles are present in the tubes and capillary of the VAST system, click on the Prime button on the LP Sampler window to remove them.
2. Consider the unsuccessful lesions (as assessed from the remaining fluorescence in the lesion site, apart from the expected residual and homogenous background), which can be due to several reasons mentioned below.
   1. Low laser power: When this happens, try with a higher value.
      NOTE: The VAST system is equipped with a dye laser. This implies that the concentration of the dye solution used for laser light generation can change with time, leading to a decrease in laser power. Replacing with a fresh solution usually solves the problem²².
   2. Poor calibration: When this happens, verify the calibration and power of the laser system as per step 5.2.2.1-5.2.2.4. If not calibrated correctly, the laser won't be directed to the desired location, thus leading to unsuccessful lesions or undesired damage in adjacent tissues.
      1. Place a mirror slide on top of the capillary chamber. Focus on the coated side (it should face the objective). Use a previous default in the slide to focus more easily.
      2. Apply a pattern of laser ablation using a calibration script.
      3. Assess the quality of the pattern. The spots or lines should appear sharp and not blurry.
      4. Use a ramp with increasing power to evaluate whether the laser power has changed compared to the previous sessions.
   3. Larval movement during lesions: Larvae respond differently to anesthesia; thus, the laser lesion may trigger movements of the tail during the process, thus preventing a successful transection. When this happens, take an extra iteration of the laser lesion steps to complete it while still avoiding damage to the surrounding tissues.
   4. Bad focus: When this occurs, focus on the middle of the central canal to get the best results.
   5. ROI drawing, position, and size: The position and size of the ROI are critical for successful transections. The ROI should be larger than the spinal cord and centered on the center of the spinal cord. To solve this, start to draw the ROI from the ventral side of the spinal cord and go up toward the dorsal side to obtain successful transection. This is likely due to tail movements triggered by the sequence of laser shots during the ablation procedure.

Results

Validation of spinal cord transection
Structural and functional investigations were performed to assess whether the protocol allows a complete spinal cord transection.

First, to verify that the loss in fluorescence at the lesion site was due to neuronal tissue damage and not fluorescence photobleaching from the laser illumination, immunostaining using an antibody against acetylated tubulin (see Table of Materials and Supplementary File 1...

Discussion

There is an urgent need for a deeper understanding of the processes at play during regeneration in zebrafish. This animal model offers many benefits for biomedical research, in particular for spinal cord injuries¹. Most of the studies involve manual lesions that require a well-trained operator and induce multi-tissue damage. A laser lesion protocol is presented here, allowing control over the lesion characteristics and reduced damage to the surrounding tissues. Furthermore, this technique is easy ...

Materials

Name	Company	Catalog Number	Comments
Software
Microscope software Zen Blue 2.0	Carl Zeiss
ImageJ/FIJI	Open-Source
Visual Studio Code	Microsoft
Microscope and accessories
ApoTome microscope	Carl Zeiss
C-Plan-Apochromat 10X (0.5NA) dipping lens	Carl Zeiss
dual AxioCam 506 m CCD cameras	Carl Zeiss
Laser scanning confocal microscope LSM880	Carl Zeiss
Spinning-disk module CSU-X1	Yokogawa
Upright microscopeAxio Examiner D1	Carl Zeiss
UV laser	Micropoint
VAST BioImager	Union Biometrica
Labware
90 mm Petri dish	Thermo-Fisher	101R20
96-well plate	Corning	3841
Chemicals
Click-It EdU Imaging Kit	Invitrogen	C10637
aminobenzoic-acid-ethyl methyl-ester (MS222)	Sigma-Aldrich	A5040
phenylthiourea (PTU)	Sigma-Aldrich	P7629
Antibodies
Donkey anti-chicken Alexa Fluor 488	Jackson	703-545-155
Donkey anti-mouse Cy3	Jackson	715-165-150
Mouse anti-GFP	Abcam	AB13970
Mouse anti-tubulin acetylated antibody	Sigma	T6793
Transgenic zebrafish lines
Tg(beta-actin:utrophin-mCherry)		N/A	Established by David Greenhald, University of Edinburgh
Tg(mnx1:gfp)		N/A	First described in [Flanagan-Steet et al. 2005]
Tg(Xla.Tubb:DsRed)		N/A	First described in [Peri and Nusslein-Volhard 2008]
Tg(Xla.Tubb:DsRed;mpeg1:GFP)		N/A	Established by Katy Reid, University of Edinburgh
Tg(Xla.Tubb:GCaMP6s)		N/A	Established by David Greenhald, University of Edinburgh