Liposomal Clodronate-mediated Macrophage Depletion in the Zebrafish Model

[Abstract] The ability to conduct in vivo macrophage-specific depletion remains an effective means to uncover functions of macrophages in a wide range of physiological contexts. Compared to the murine model, zebrafish offer superior imaging capabilities due to their optical transparency starting from a single-cell stage to throughout larval development. These qualities become important for in vivo cell specific depletions so that the elimination of the targeted cells can be tracked and validated in real time through microscopy. Multiple methods to deplete macrophages in zebrafish are available, including genetic (such as an irf8 knockout), chemogenetic (such as the nitroreductase/metronidazole system), and toxin-based depletions (such as using clodronate liposomes). The use of clodronate-containing liposomes to induce macrophage apoptosis after phagocytosing the liposomes is effective in depleting macrophages as well as testing their ability to phagocytose. Here we describe a detailed protocol for the systemic depletion of macrophages in zebrafish larvae by intravenous injection of liposomal clodronate supplemented with fluorescent dextran conjugates. Co-injection with the fluorescent dextran allows tracking of macrophage depletion in real time starting with verifying the successful intravenous injection to macrophage uptake of molecules and their eventual death. To verify a high degree of macrophage depletion, the level of brain macrophage (microglia) elimination can be determined by a rapid neutral red vital dye staining when clodronate injection is performed at early larval stages.

[Background] Macrophages are key constituents of the innate immune system and play important roles in response to infections, sterile inflammation, and environmental changes. One of the most effective ways to uncouple the functions of macrophages from a complex mix of interacting cell types in different physiological contexts is to be able to specifically eliminate macrophages and analyze the phenotypic consequence. Such depletion experiments in mice have provided much insight into the role of macrophages (Hua et al., 2018;Rosowski, 2020). However, our understanding of macrophage functions remains incomplete, and cell depletion experiments in the mouse model are difficult to track and validate in real time. For these reasons, the optical transparency and ease of manipulation of larval zebrafish offer clear advantages for highly traceable and tractable cell ablation in vivo by imaging the target cells in real time and in the whole intact organism. Zebrafish genes and immune system also share a high degree of orthology with those of the human (Yoder et al., 2002;Santoriello et al., 2012;Howe et al., 2013). Furthermore, the adaptive immune system of zebrafish does not become functionally mature until juvenile adult stages (Lam et al., 2004), making the larval zebrafish an excellent platform to study the innate immune system independent of the adaptive immune contributions.
Currently available macrophage depletion methods in zebrafish include genetic and chemogenetic manipulations, and toxin-based depletion. The development of macrophages requires early and continual function of the transcription factor Pu.1 (with the gene name spi1b), together with another transcription factor Irf8 (Li et al., 2011;Shiau et al., 2015;Tenor et al., 2015). Either disruption of pu.1 or irf8 by gene knockout or knockdown by morpholino (MO) anti-sense oligomers, makes a reliable 3 www.bio-protocol.org/e3951 approach for macrophage depletion, while the former ablates myeloid cells, and the latter is more specific to macrophages but also causes an increase in neutrophil numbers (Shiau et al., 2015;Yang et al., 2020). These methods are not amenable to temporal control (Rhodes et al., 2005;Li et al., 2011;Shiau et al., 2015;Rosowski, 2020), while clodronate-mediated depletion of macrophages based on localized microinjection enables some degree of spatial and temporal designation (Bernut et al., 2014).
Clodronate (also known as dichloromethylene diphosphonate) can be metabolized by cells to block mitochondrial respiration due to the formation of a non-hydrolyzable ATP analog, which then causes cell death (apoptosis) (Rosowski, 2020). Once injected as encapsulated in liposomes, clodronate is easily ingested and eliminated by macrophages as it accumulates within the cell (van Rooijen and Hendrikx, 2010). Since neither the clodronate nor the liposomal phospholipids utilized are toxic to other nonphagocytic cells (van Rooijen and Hendrikx, 2010), this approach allows specific depletion of already existing phagocytosing macrophages.
As part of the design for our protocol, we co-injected fluorescently labeled dextran with the liposomal clodronate to allow us to verify a precise and accurate injection and track the effect of the clodronate on macrophages in the entire larvae. To this end, after intravenous co-injection of clodronate liposome with fluorescently tagged dextran, we visually verified the successful injection of the substances into time we found that our injected substances easily reach macrophages throughout the body including the brain. Using the 48-hour window for clodronate to take effect, we were able to achieve full ablation of microglia in most injected larval zebrafish (Yang et al., 2020). Overall, we found that intravenous microinjection of liposomal clodronate at 3 dpf with a 48-hour incubation time is effective in eliminating macrophages.   Microelectrode puller program (see Table 1)  b. The goal is to have the larvae embedded in a thin layer of agarose so that when it comes to using the fine glass needle for injection, the needle does not need to penetrate through much gel to reach your desired tissue target (see Figure 2). Too much agarose will make it hard to direct the needle to your target site without having it bend or swerve in the agarose. 4. The most time-sensitive step is in orienting all the zebrafish larvae quickly in the agarose before it begins to re-solidify. You can expect to have less than 1-2 min for this step. Use fine forceps to quickly orient each one on their sides, but without concern of the exact orientation or body alignment, only that the tail region containing the caudal vein plexus is accessible to your needle.

Materials and Reagents
Take caution to not poke the larvae to cause injury but to use the agarose around the larvae to move them. 5. Wait for agarose around the larvae to cool down and solidify before starting injections. 6. Add a small amount of fish system water to cover the agarose to prevent dehydration and keep the larvae healthy during the operation (see Figure 2). E. Intravenous microinjections 1. Use the foot pedal to inject 1 nl of clodronate liposomes or control liposomes into caudal vein plexus, which is located in the tail tissue right past the yolk extension (see Graphical abstract).
2. Validate each injection by seeing an immediate distribution of the injected fluorescent dextran mix into the heart and subsequent circulation throughout the body vasculature using a fluorescent stereomicroscope. Any larvae after injection that do not show the correct distribution of the fluorescent marker in the body vasculature is removed from the dish using forceps.
3. Gently recover the injected fish from the agarose into fresh fish water supplemented with 0.003% PTU and transfer them back into the 28.5 °C incubator.
Use the side of the forceps to create a break in the agarose beginning underneath the head and dragging the forceps along the body until the tip of the tail is reached. Larvae should be able to squirm out easily and swim into the system water surrounding the agarose. Pipette released larval zebrafish into the dish with fresh fish water. 4. After microinjection, monitor all the fish for normal health and behavior. Remove any sickly or abnormal fish, and only continue processing the healthy and normal larvae.
F. Neutral red staining 1. In a Petri dish, stain a subset of your injected larvae at 48 hpi (hours post injection) with neutral red at 2.5 μg/ml in fish water supplemented with 0.003% PTU at room temperature for 45 min to 1 h (see Figure 3A).
2. Remove the majority of the neutral red supplemented fish water by exchanging it for fresh new fish water supplemented with PTU (see Figure 3B).
3. Leave the larvae in incubator for another 2-3 h before analyzing under a stereomicroscope.
4. Before analyzing, anesthetize larvae with tricaine at 0.5× strength, which the larvae can stay in for up to 45 min with the ability to fully recover back to normal and moving again. For the anesthetics to be working, larvae should not be moving so you can use them for your analysis.
Note: Please follow your institutional guidelines on the appropriate protocol for anesthetizing zebrafish.   Store concentrated stock at -20 °C. To use tricaine as an anesthetic ， use at a 0.5-1.0× concentration.

3% Methyl cellulose
Note: You will need 3 g of methyl cellulose powder in 100 ml of distilled water.
a. To dissolve methyl cellulose powder in water, bring half of the distilled water to 60 °C. The remaining volume of water should be stored at 4 °C and left to cool for 30 min. b. Add 3 g of methyl cellulose powder to boiling water and stir. The mixture will become extremely viscous quickly.
c. Remove the solution from the heat as soon as stirring becomes difficult. d. Add the chilled water and continue to stir for 30 s. e. To fully dissolve particles and remove bubbles the solution must be placed at -20 °C overnight. To prevent glass from breaking, gradually lower the temperature by placing glass at 4 °C for 1-2 h before placing at -20 °C overnight.

1,000× neutral red dye solution
Dissolve neutral red powder in distilled water at a concentration of 2.5 mg/ml for a 1,000× stock.
For live staining of microglia in zebrafish, use neutral red at 1× strength.
5. 1.5% low melt agarose a. Dissolve 1.5 g of low melt agarose in 100 ml distilled water using the microwave for less than 30 s. b. Monitor the heating process as low melt agarose can easily overheat and bubble over.
c. Once this solution is made, it can be stored in a glass bottle with a lid at room temperature and be repeatedly re-used by re-melting the agarose in the microwave.