Automating Tumor Implantation in Zebrafish Larvae for Cancer Research and Medicine

Summary

Here, we present a detailed protocol for automated tumor xenograft implantation into zebrafish larvae using a state-of-the-art robotic injection system. Furthermore, we present injection videos and validation data demonstrating the system's successful performance.

Abstract

Zebrafish larval injections, including subcutaneous and brain-orthotopic tumor xenografting as well as intravenous drug administration, have become central techniques in zebrafish-based research. These procedures, however, remain tedious, technically challenging, and highly dependent on operator skill, limiting throughput and reproducibility. To address these limitations, we introduce an advanced robotic system that automates microinjections into zebrafish larvae, significantly improving efficiency, precision, and reproducibility over manual injections. Here, we provide a detailed protocol for operating the automated injector, a high-throughput system capable of delivering liquids, particles, or cancer cells at three different anatomical sites with superior accuracy and speed compared to trained zebrafish researchers. We outline the essential steps, including setup, calibrations, injection procedures, and troubleshooting, to ensure optimal performance. Furthermore, we provide representative examples of results from using the robot and discuss its potential to facilitate large-scale larval injections, including for drug screens and precision medicine. By integrating an automated injector into the workflows, researchers can overcome limitations of manual injections and accelerate preclinical drug discovery and personalized treatment selection.

Introduction

As our understanding of cancer behavior and drug resistance improves, it has become increasingly clear that there is a need for better diagnosis, characterization, and therapeutic targeting of cancer subtypes. During the last twenty years, publications in the field of personalized medicine (also known as precision medicine) have been on the rise^1,2. Indeed, precision medicine is defined as the practice of tailoring medical treatment based on the individual patient's disease characteristics³. This follows our current understanding that there is no one-drug-fits-all approach, which has become particularly clear and pertinent in cancer therapy. Currently, however, the lack of clinically relevant precision medicine approaches for most cancer patients means that the process of selecting the most suitable treatment strategy is by trial and error. This results in significant delays and unnecessary discomfort with diminished quality of life and shortened survival. Importantly, a patient with diminished quality of life after several ineffective chemotherapy regimens may opt to switch to palliative treatment⁴. With a suitable, personalized screening method, drug selection can be performed with higher precision and without delay. While it is possible to perform genetic and proteomic sequencing on cancer cells⁵, such molecular data is overall poorly predictive of patient outcome and often has little clinical value⁶. Within drug development, having a precision medicine approach as early as possible greatly improves success rates at the various milestones of the drug development pipeline. Indeed, understanding early on which drug candidates have the broadest inter-individual efficacy profile, or what cancer subtypes are particularly sensitive to each drug candidate, de-risks and improves the efficiency of late preclinical tests and clinical trials⁷. With rising pharmaceutical R&D costs⁸, growing political pressure to lower drug prices^9,10, and advances in AI-driven target validation¹¹, the need for a large-scale, efficient, and precision-medicine-centered screening method in drug development has never been greater.

Here, zebrafish enter the picture, specifically zebrafish larvae at 48 hour post-fertilization (hpf). Tumor xenograft models based on zebrafish larvae as hosts constitute a rapid in vivo screening platform to forecast the clinical responses to chemotherapy^6,12,13,14. Zebrafish tumor xenograft models have emerged as a powerful alternative to both molecular precision medicine (i.e., genomics or molecular pathology approaches to guide choice of treatment for cancer patients), as well as organoid and mouse xenograft studies for preclinical drug development^15,16. With zebrafish tumor xenografts, fluorescent tumor cells (genetically or chemically labelled) are engrafted into zebrafish larvae with high efficiency¹³. Drugs or drug candidates with a relatively fast mode of action (e.g., such that target tumor cell viability directly or indirectly via, for example, activation of T-cell cytotoxicity) can then be screened in just three days, leading to fast, functional readouts of anti-cancer and anti-metastatic drug efficacy. Following this, an informed selection of the therapy that shows a good efficacy can be made, with studies showing that this should translate to a strong treatment outcome for the patient or a chance for success in a drug development project respectively^12,17,18,19. Moreover, this screening process is highly relevant for selecting compounds to be moved forward in the drug development pipeline.

However, to become a viable diagnostic tool in clinical settings and to enable large-scale drug screens within cancer research and drug development, the zebrafish tumor xenograft (ZTX) model must be fast, reliable, reproducible, and cost-effective. Automation is essential for achieving these goals, as it minimizes operator variability while increasing throughput and enhancing precision in tumor cell injection and analysis. By standardizing these procedures, we can improve data reproducibility and eliminate human error, while the automated workflows present opportunities for high-throughput screening and making large-scale studies feasible^20,21. Recently, we have developed an automated injector that injects liquids, particles, or cells of various origins at three different anatomical sites with equal or higher speed, precision, and reproducibility than trained zebrafish researchers²². Here, we present a protocol for the automated injection of tumor cells into zebrafish larvae in a precise and controlled manner. Using advanced image recognition, zebrafish larvae are located on the injecting plate, orientation is determined, and the target site for the injection is identified. The needle is then guided automatically, and a successful puncture is determined by the software. With minimal training, operators can use this system to perform complex injections, for example, creating microtumors in live zebrafish larvae, thus enabling efficient screening of treatment strategies.

Protocol

Zebrafish lines were maintained according to standard protocols (https://zfin.org) and handled in compliance with local animal welfare regulations. These regulations implement the Council of Europe's Guidelines for the Protection of Experimental Animals (Directive 86/609/EEC), which permit the use of zebrafish larvae up to the onset of free feeding (5 days post-fertilization). As all larvae used in this study were no more than 5 days old, no ethical license was required under Directive 86/609/EEC.

NOTE: Dechorionated 48 hpf zebrafish larvae and the material to be injected in these larvae should be prepared prior to starting the procedure using the automated injector. Protocols for this can be found in the supplemental methods (Supplementary File 1) associated with this article. A schematic presentation of the protocol workflow is shown in Figure 1. Supplementary Figure 1 shows a schematic representation of the developmental stages of zebrafish embryos.

1. Microneedle preparation

NOTE: Microneedles are required for the injections and are an essential element of the injection process to troubleshoot and tune for this purpose. Both commercial needles and manually made needles can be used in the automated injector. The following is the procedure for manually pulling microneedles using Sutter Instrument P-1000 or PC-10, although most microneedle pullers are similar in operation. The preferred needle morphology is a gradual 6-8 mm taper, with a tip size of 0.3-0.7 µm, which is manually broken open before injection using sharp forceps.

1. Prepare suitable borosilicate glass capillaries: 1.0 mm OD, 0.78 mm inner diameter (ID). Ensure that they are clean and free from dust and debris.
2. Check that the heating coil is intact and in good condition.
3. Insert the glass capillary into the clamps and secure firmly using the clamp screws but not so tightly that the tube is cracked.
4. Adjust the settings: Heat setting, pull strength, and ramp time. When using a PC-10 needle puller, the correct needle morphology is obtained using a single-stage pull with the heating setting 68 and full weights.
   NOTE: These settings will influence the length and width of the tip taper and may vary depending on the wall thickness of the capillary. Trial and error are required here to produce the best result.
5. Remove the needles from the clamps carefully, not to damage the tip. Inspect the tip to ensure an even taper.
6. Store the microneedles carefully on adhesive tape, ensuring that the delicate tip does not contact any surface.

2. Agarose gel plate preparation for robotic injection

1. Weigh 0.6 g of agarose and transfer it to a 100 mL glass bottle.
2. Using a measuring cylinder, measure 40 mL of E3 solution. Place it into the glass bottle with the agarose to prepare a 1.5% agarose solution.
3. Place the glass bottle in a microwave and heat for 30 s at 900 W. Then, continue heating in 30-s increments, swirling the contents (wearing gloves) between each interval until the agarose is completely dissolved.
   NOTE: Be careful, as boiling can also start as a result of swirling.
4. Once melted, cool the agarose by flushing the exterior of the glass bottle under running water until it feels warm but not hot to the touch (approximately 60 °C).
5. Clean and thoroughly dry the reusable plate holder and glass slide. Assemble by sliding the glass slide into the slot on the side of the plate holder (Supplementary Figure 2A) and place it on a flat surface.
   NOTE: Check that the glass plate is not sticking out of the side of the holder.
6. Pour approximately 40 mL of the agarose mixture onto the glass plate, ensuring it spreads to cover the entire surface. If necessary, tilt around such that all edges have been reached.
7. Allow the agarose to solidify for about 5 min. The plate is then ready for use.
   NOTE: Adding E3 solution on the surface of the solid agarose plate can prevent drying. However, it should be removed prior to its use for injections.

3. Placing zebrafish larvae on the agarose gel plate

1. Moisten the agarose gel plate with E3 solution.
2. Hold the plate at an angle, allowing excess E3 solution to be collected.
3. Prepare an E3 solution containing 40 µg/mL tricaine in a Petri dish. Place approximately 20 larvae into the solution and allow them to anesthetize for 2 min.
4. Carefully place 10-20 anesthetized larvae onto the plate (Supplementary Figure 2B), ensuring that excess water drains away naturally.
5. Make sure that the larvae are spread out, not touching one another, and avoid positioning them too close (5-10 mm) to the edge of the plate (Supplementary Figure 2B).
6. Remove any excess liquid from the plate while ensuring that the larvae remain moist.
   NOTE: Excessive liquid can induce unintended larval movement during injection, while inadequate moisture levels may increase mortality rates.

4. Loading the microneedle

NOTE: Load the microneedle with high-density, fluorescent cells just prior to performing an injection. Cells can genetically express a fluorescent marker, or a chemical label can be used²³. Although the number of cells injected in each larva is comparatively low, typically, 700 to 900 cells, the cell density needs to be high at the injection stage; around 300,000 to 500,000 cells per µL, due mainly to the minute volume (around 2 nL) that is injected.

1. Centrifuge the cell suspension at 200 × g for 5 min and remove the cell media.
2. Count the cells using a cell counter, then resuspend the pellet in sterile, room-temperature (RT) PBS to achieve a final density of 300,000 cells/µL.
3. Take a prepared microneedle and hold it carefully between the thumb and forefinger. Then, using a microloader pipette tip, load the microneedle with 1-10 µL of the cell suspension, depending on the number of cells.
   NOTE: Insert the tip to the end of the microneedle and slowly draw it out while it is being filled.
4. Using a stereomicroscope at 4x magnification and a pair of fine forceps, carefully break off the sealed tip of the microneedle to produce an opening. This step can also be performed prior to loading an injection suspension.
   NOTE: A beveled needle tip facilitates puncture and can be created using a needle grinder. If using a needle grinder, bevel the needle tip before loading the injection suspensions. In this case, the needle is already open before cells are loaded.

5. Operation of the robot for microinjection

1. Perform the booting procedure.
   1. Ensure an adequate gas supply is connected and turned on.
      NOTE: The recommended input pressure from an air compressor or an air valve in a laboratory setting with centralized, pressurized air is 0.6 MPa (6 bar). The air compressor or valve can remain continuously on and open without the need for frequent switching off. Compressed air, nitrogen, or CO₂ can be used for the input pressure.
   2. Insert a USB drive into one of the ports at the back of the robot (optional).
      NOTE: A USB drive should be inserted into the machine prior to powering up the robot and is removed only after the robot has been powered off.
   3. Press the green power button next to the screen to turn on the robot (Supplementary Figure 2C). Wait till the control panel shows up (approximately 1 min).
   4. Check if there is 15 cm of free space on the left and right sides of the robot and press OK on the popup (Supplementary Figure 2D).
   5. Press OK on the next popup (Supplementary Figure 2E) about the USB drive.
      NOTE: This popup will not appear if no USB drive is inserted.
2. Set the connection for remote control (optional).
   NOTE: A Remote control is used to view and operate the machine from a distance. This function is useful for demonstrations, training sessions, software updates, and troubleshooting.
   1. Insert an Ethernet cable into the port at the back of the machine. For initial setup, contact the IT department for assistance.
   2. Tap the menu icon located at the top-left corner of the screen (Supplementary Figure 2F).
   3. Tap the settings icon at the bottom center of the menu (Supplementary Figure 2F).
   4. Select Maintenance mode from the settings menu (Supplementary Figure 2G).
   5. Tap Connect to LSM to initiate a remote connection (Supplementary Figure 2H).
3. Perform injection settings.
   1. Tap on injection settings in the menu (Supplementary Figure 2F).
   2. Choose the appropriate developmental stage of zebrafish larvae for injection (Supplementary Figure 3A).
      NOTE: For injections to the duct of Cuvier, perivitelline space, and hindbrain ventricle, 48 hpf larvae kept at 28.5 °C are recommended.
   3. Select the preferred injection site from the options provided for performing injections (Supplementary Figure 3B).
   4. Select the injection location on the schematic diagram of a zebrafish larva. Adjust the position of the larval diagram, needle tip, and needle orientation to align with the selected injection site (Supplementary Figure 3C).
      NOTE: Suggested needle placements for different injection sites are provided in a previously published paper²². For the Duct of Cuvier (DoC) injection, typically, the needle is placed in the middle of the DoC due to the higher blood flow in that region. The needle orientation can be either dorsal or ventral. For perivitelline space (PVS) injection, it is recommended to begin the needle insertion at the border of the space and position it at approximately a 30-45° angle to the notochord. For hindbrain ventricle injection, it is recommended to commence the needle insertion outside of the hindbrain to prevent it from slipping. The needle angle should align between the eyes and the otic vesicle.
   5. Select the injection macro, offering options for both automatic and manual modes (Supplementary Figure 3D).
      NOTE: For automatic mode, the name of the injection site indicates that the injection process is performed autonomously by the system. Manual selection entails the robot autonomously locating and approaching the larvae. Once the larvae are identified, a control interface will appear for users to operate the injection process.
   6. Once the injection settings are finalized, tap on Continue to needle calibration to proceed with calibrating the needle.
4. Perform needle calibration.
   NOTE: As each needle is different, the needle tip position needs to be calibrated. The initial interface of needle calibration is shown in Supplementary Figure 3E.
   1. Grasp the needle holder firmly and rotate the metal part counterclockwise to detach the needle holder from the robot.
   2. Insert the blunt side of a filled needle into the frustoconical side of the needle holder.
   3. Tap on Move stage to mount needle displayed on the screen (Supplementary Figure 3E). The stage will relocate to a secure position for mounting the needle, and a ruler will be displayed (Supplementary Figure 3F). Measure the length of the needle outside of the needle holder using the ruler. It should be around 22 mm.
   4. Grasp the needle holder containing the filled needle and twist the metal part clockwise to securely tighten the connection (Supplementary Figure 3G). Press OK to proceed.
   5. In the popup screen, position the needle at the center of the circle (Supplementary Figure 3H) by adjusting the two screws on the machine (Supplementary Figure 3I). When centered, tap the green checkmark.
   6. In the next popup screen (Supplementary Figure 3J), adjust the focus on the needle tip by turning the screw located at the back of the robot head (Supplementary Figure 3K). When focused, tap the green checkmark. The robot will then automatically calibrate the needle height. Subsequently, the robot will rotate the needle and display the injection positions as red points with each rotation (Supplementary Figure 3L).
   7. When finished, tap Got it and Continue to droplet calibration at the bottom right of the screen.
5. Perform droplet calibration.
   NOTE: The droplet size is calibrated using an injection in mineral oil.
   1. Prepare a 6-well plate with one well filled with 10 mL of mineral oil and another well with 10 mL of cleaning buffer (e.g., PBS or water). Place the 6-well plate on the left position of the plate holder.
      ​NOTE: Please remove the lid of the 6-well plate during the (whole) experiment so it can be used again when a needle is clogged or a new needle is placed.
   2. Tap the icon next to Calibrate using a well plate (Supplementary Figure 4A).
   3. Select the positions of the oil well and cleaning well (Supplementary Figure 4B).
   4. Tap on Make droplet to generate a droplet with the settings shown on the left side of the screen (Supplementary Figure 4C). Adjust the injection pressure, back-pressure, and air pulse duration as needed to reach the target volume. Alternatively, tap the Auto Improve button to automatically optimize the settings and achieve the desired volume.
   5. If the needle is clogged, press the blue Clean needle button (Supplementary Figure 4C) to activate a high-pressure cleaning pulse in the well containing water or medium.
   6. If the needle and droplet appear blurry, adjust the focus of the needle and droplet using the top third button on the right side of the screen (Supplementary Figure 4C).
   7. Tap on the green checkmark to close the screen.
6. Perform plate selection.
   1. Click Move stage to place plate, and the robot will automatically move the stage to an appropriate position for placing the plate.
   2. Check that the agarose gel plate is dry underneath and place the plate with the anesthetized larvae in the right position of the plate holder.
   3. Then, click OK and tap on Continue to injection to proceed with the injection process (Supplementary Figure 4D).
7. Perform injection.
   1. In the injection interface, click the Start button to begin the injection process (Supplementary Figure 4E). The process begins with the robot stage moving while the system scans the agarose plate, searching for a fish (Supplementary Figure 4F). Upon identifying a larva, the needle automatically navigates to the designated injection site and automatically adjusts its direction to the predetermined angle.
   2. If automatic mode is chosen, the robot seamlessly administers injections at the selected site (Supplementary Figure 4G). Upon completing the current injection, the robot will continue scanning for the next larva on the plate to do injections.
   3. If manual mode is selected, a manual control interface will appear when the needle approaches the larva (Supplementary Figure 4H). Use the blue and yellow arrows to move the needle, and rotate the needle using the circular rotation button on the left side of the screen. Once the needle is positioned inside the larva, tap the red droplet button to perform the injection. Alternatively, tap the m key to initiate the macro sequence to perform injection automatically.
   4. When all larvae have been injected, close the injection interface by clicking the stop button. The needle will be placed in the oil well automatically.
   5. Carefully remove the plate containing the injected larvae.
8. Transfer the injected zebrafish larvae.
   1. To transfer the injected larvae, tilt the plate over a Petri dish and use a pipette or squeeze bottle to gently flush out the larvae with a controlled flow of water.
   2. Start from the top and move downwards for effective flushing.
   3. Remove any dead larvae, replace the water with fresh E3 solution, and place the Petri dish in the incubator.
      NOTE: If human cancer cells or clinical samples are injected, incubate the Petri dish at 33-36 °C; otherwise, incubate at 28.5 °C.

6. Switching off the robot

1. When the injection is complete, remove the needle by visiting the needle calibration menu and pressing Move stage to remove needle (Supplementary Figure 3E). If one intends to keep the needle in the machine for later use, skip this step.
2. Go to the menu and tap on the power button, located at the bottom left (Supplementary Figure 2D).
3. Press Park needle button to preserve the needle (Supplementary Figure 4I).
4. Otherwise, click OK. The machine will shut down, and the green power button light will turn off once the shutdown is complete.
   NOTE: When the robot is idle, place empty multi-well plates in both multi-well plate holders to prevent dust from entering the machine.

7. Removing agarose gel and clean up

1. Remove the agarose gel from the plate and carefully (wear goggles and gloves) remove the glass plate from the holder.
2. Clean the glass plate and the holder thoroughly with warm water and ethanol, ensuring it is dried before reuse.
   NOTE: Clean all gel plates immediately after use, as agarose gel is easier to remove while still wet.

Results

Here, we present the automated injection process for the three most used injection sites for cancer cell xenografts.

Video 1 demonstrates the automatic injection of the duct of Cuvier (DoC) using the robotic system. The robot first scans the plate systematically, moving from top to bottom and left to right, to locate a zebrafish larva. Once a larva is identified, the needle automatically approaches it, adjusting to the predetermined starting position and needle angle based on ...

Discussion

The development and implementation of the automated injector represent a significant advancement in the field of zebrafish tumor xenografting for anti-cancer drug screening and precision medicine. This is, to the best of our knowledge, the first commercially available robot for automated injection of cell suspensions into zebrafish larvae, and this protocol is therefore relevant for all research that involves such injections in high throughput. By automating the injection process, this system furthermore minimizes operat...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
6-well plate	Greiner bio-one cellstar	ref 657160
Agarose	VWR	#N605-100G
Automated injection system + consumables	Life Science Methods	https://www.lifesciencemethods.com/microinjection_system_for_zebrafish_larvae.html?nocache=1752058717774
Borosilicate glass capillaries	World Precision Instruments	N/A	1.0 mm OD, 0.78 mm ID
Cell counter	ThermoFisher	Countess 3
Commercial needles	Clunbury Scientific LLC	#B100-58-20
DNase type I	Roche	#11284932001
Filter for cancer cells	Sysmex CellTricsTM	04-0042-2316	30 µm
Mineral oil	Diapharma	#330779-1L
Needle grinder ()	Narishige	EG-401
Polyvinylpyrrolidone 40	Sigma	#102420477
GentleMACS Octo Dissociator	Miltenyi Biotec	https://www.miltenyibiotec.com/US-en/products/gentlemacs-octo-dissociator-with-heaters.html