A zWEDGI Technique to Visualize Fungal Pathogen-Infected Zebrafish Larvae

1. Preparation of Aspergillus spores for injection

1. From an Aspergillus spore suspension, calculate the volume needed to obtain 1 x 10⁶ spores. The volume should be 20–100 μL; if not, produce a 10x dilution in 0.01% (v/v) sterile Tween-20 (Tween-water; Table of Materials). For example, if the calculated volume is 5 µL, produce a 10x dilution and use 50 µL of the diluted solution.
   NOTE: Two plates/strains can be prepared to collect more spores or as a spare in the case of contamination.
2. Spread 1 x 10⁶ Aspergillus spores on one glucose minimal media (GMM) plate (Table of Materials) with a sterile disposable L-shaped spreader in a biosafety cabinet. Avoid spreading to the margin of the plate. Incubate at 37 °C for 3–4 days, with the plate facing upside down.
3. On the day of collection, bring sterile miracloth and 50 mL conical tubes (two per strain), fresh bottles of sterile Tween-water (one per strain), and sterile disposable L-shaped spreaders to the biosafety cabinet.   
   NOTE: Miracloth can be cut into ~8 in x 6 in pieces, wrapped in foil, and autoclaved to sterilize.
4. Place one piece of miracloth in each labeled 50 mL conical tube and re-cap. Take the remaining miracloth packet out of the hood.
5. Bring plates into the biosafety cabinet. Open one plate, then pour Tween-water on the top to cover about three-quarters of the plate.
6. Using a disposable L-shaped spreader, gently scrape the surface of the fungal culture in a back-and-forth motion, while using the other hand to rotate the plate. Scrape until almost all of the spores are homogenized into the Tween water.
   NOTE: Due to high hydrophobicity, spores can create "puffs" when Tween water is added or during scraping. Great care should be taken to avoid contamination of nearby tubes or plates. It is advised to change gloves and wipe off the surface with 70% ethanol between the extraction of different strains.
7. Take one 50 mL conical tube and remove the piece of miracloth. Fold it in half and make it into a filter inserted in the top of the 50 mL conical tube.
8. Pour the fungal homogenate from the plate over the miracloth into the tube.
   NOTE: If two plates of one strain are prepared, scrape both plates and pour them into the same conical tube.
9. Pour Tween water to bring the total volume in the conical tube to 50 mL.
10. Spin at 900 x g for 10 min. Make sure to use aerosolization-preventing caps in the centrifuge.
11. Pour off the supernatant into ~10% bleach solution to decontaminate. Pour 50 mL of sterile 1x phosphate-buffered saline (PBS) into the conical tube, then vortex or shake to resuspend the pellet.
12. Spin again at 900 x g for 10 min. Pour off the supernatant and resuspend the pellet in 5 mL of sterile 1x PBS. Filter through a fresh piece of miracloth into a fresh 50 mL conical tube.
13. Make 10-fold serial dilutions (10x, 100x, 1000x) of the fungal homogenate in 1.7 mL centrifuge tubes (e.g., for the 10x solution, mix 100 µL of the fungal homogenate with 900 µL of Tween-water).
14. Choose the first dilution in which the spores are not visible when it is discharged into Tween-water and use this dilution to count the number of spores using a hemocytometer.
15. Calculate the spore concentration in the prepared fungal homogenate (water suspension) using the following formula:
    Concentration (spores/mL) = Number of spores in middle 25 boxes x dilution factor x 10⁴
16. Prepare a 1 mL stock of 1.5 x 10⁸ spores/mL in sterile 1x PBS in a 1.7 mL microcentrifuge tube. This spore preparation can be stored at 4 °C for ~4 weeks.
17. Prior to use in injections, mix 20 µL of the spore preparation with 10 µL of 1% sterile phenol red in a 1.7 mL centrifuge tube to achieve a final spore concentration of 1 x 10⁸ spores/mL. Vortex thoroughly prior to injection.        
    NOTE: 1% phenol red solution should be filter-sterilized and stored in aliquots.
18. For a mock injection, mix 20 µL of 1x PBS with 10 µL of 1% sterile phenol red.

2. Preparation of agar plates for injection

1. Prepare 2% agarose in E3 medium and melt in a microwave.
2. Pour into a 100 mm x 15 mm Petri dish (~25 mL per plate), swirl to cover the plate evenly, and let cool.
3. Wrap the plate with paraffin film and store inverted at 4 °C.
4. Prior to injection, bring the plate to room temperature (RT).
5. Pour ~1 mL of filter-sterilized 2% bovine serum albumin (BSA) onto the plate, tilt the plate to spread and cover the entire bottom, and rinse with E3.
   NOTE: 2% BSA solution can be filter-sterilized and stored as 1 mL aliquots at -20 °C. 2% BSA pre-treatment prevents larvae from sticking to the surface of the agarose.
6. Pour E3 with buffered tricaine onto the plate and let it sit until injection.

3. Zebrafish larva hindbrain ventricle microinjection

1. Manually dechorionate larvae with forceps at 2 dpf in a Petri dish.
   NOTE: Dechorionation can be performed anytime from 1.5 dpf until the time of injection.
2. Remove as much E3 as possible from the Petri dish and add buffered 300 µg/mL tricaine in E3 to anesthetize larvae.       
   NOTE: A stock solution of buffered 4 mg/mL tricaine in E3 can be prepared and stored at 4 ˚C. The working solution can be made by diluting 4 mL of the stock solution up to 50 mL with E3.
3. Use a microinjection setup supplied with the pressure injector, back pressure unit, footswitch, micropipette holder, micromanipulator, and a magnetic stand and plate, all connected to a source of compressed air (Table of Materials).
4. Open the compressed air valve and turn on the microinjector. Set the pressure to ~25 PSI, pulse duration to 60 ms, and back pressure unit to 1 Pound per square inch (PSI).
5. Load a microinjection needle using a microloader pipette tip (Table of Materials) with about 3–5 µL of prepared PBS or spore suspension with phenol red. Mount the needle onto the micromanipulator.   
   NOTE: Microinjection needles can be prepared as described previously. The stereo microscope used for microinjections should have an eyepiece reticle to calibrate the microinjection needle. The reticle should be calibrated with a stage micrometer, and the length of the reticle scale (µm) should be determined. The diameter of the spore suspension drop that ejects from the needle is measured depending on the number of hashes (of the reticle) that overlap with the drop.
6. Position the micromanipulator so that the end of the needle is in view at the lowest magnification under the stereo microscope. Zoom to 4x magnification, keeping the needle in view.
7. Using sharp forceps, clip the end of the needle. Press the injection pedal to visualize the size of the droplet that comes out. Keep clipping back until ~3 nL of spore suspension is injected (here, this is five hashes).
8. Move the micromanipulator and needle out of the way to avoid accidentally hitting the needle while the larvae are arranged on the injection plate.
9. Pour E3-Tricaine off the injection plate, then transfer ~24 anesthetized larvae to the injection plate with as little E3 as possible using a transfer pipet.
10. Using a small tool for manipulating zebrafish larvae (i.e., hair loop tool or eyelash tool), arrange the larvae according to the direction in which they are facing. Specifically, place all facing to the right in one row, and all facing to the left in a row below.    
    NOTE: This arrangement is difficult if there is too much liquid on the plate, as the larvae will "float" out of place. However, too little liquid is also problematic if the injections take a long time, as the larvae can dry out or anesthesia wear off. Thus, careful attention should be paid to the amount of liquid on the plate throughout the entire microinjection process.
11. Adjust the microscope zoom to the lowest magnification. Bring the micromanipulator back and arrange it so that the needle is close to the larvae, at a ~30˚–60˚ angle, in the middle of the field of view.
12. Zoom in to the highest magnification and use fine adjustment knobs to further adjust the position of the needle. Verify that ~30–70 spores are coming out of the needle by injecting the spore suspension into the liquid on the plate next to the larvae. Adjust the time and pressure on the injection setup, if necessary.
    NOTE: This test should be repeated every five to six larvae, as the number of spores coming out of the needle can increase or decrease over time.
13. Starting with the row in which the larvae are facing towards the needle, move the plate so that the needle is directly above and positioned near the first larvae.
14. Moving the needle with the micromanipulator, insert the needle through the tissue around the otic vesicle to pierce through into the hindbrain ventricle. Move the plate with the other hand as necessary to get the right orientation of the larva with the angle of the needle.
15. Visually verify that the end of the needle is in the center of the hindbrain ventricle, press the foot pedal to inject spores, and gently retract the needle.  
    NOTE: The phenol red dye should primarily stay within the hindbrain ventricle. A small amount may go into the midbrain, but it should not reach the forebrain or outside the brain. If it does, the volume being injected is too large, and the pressure and time should be decreased accordingly, or a new needle should be calibrated.
16. Moving down the plate, inject all the larvae in that row. Then, turn the plate around and inject all larvae into the other row.        
    NOTE: Unsuccessfully injected or accidentally damaged larvae can be marked by 1) injecting into the yolk a couple of times to create a red mark or 2) dragging the larva out of the row with the needle.
17. Move the needle up and out of the way again. Zoom out to a lower magnification on the microscope. The phenol red dye should still be visible in the hindbrain of each larva.
18. First, pull away with a hair loop tool and pipette up to dispose of any larvae with unsuccessful injections. Transfer the remaining larvae into a new Petri dish by washing them off the plate with fresh sterile E3 and a transfer pipet.
19. Repeat as necessary for the final experimental sample number desired.
20. Rinse larvae at least 2x with E3 and ensure recovery from anesthesia.
21. To quantify survival without any further treatment, using a transfer pipette, transfer larvae into a 96-well plate (1 larva per well) in E3.

4. ​Establishment of injected and viable spore numbers

1. Immediately after injection, using a transfer pipette, randomly pick about eight of the injected larvae and transfer them to 1.7 mL centrifuge tubes (one larva per tube).
2. Euthanize larvae with tricaine or by placing them at 4 °C for 0.5–2.0 h.
3. Prepare 1 mL of 1 mg/mL ampicillin and 0.5 mg/mL kanamycin antibiotic solutions in sterile 1x PBS. The leftover solution can be stored at 4 °C and used later.
   NOTE: Stock solutions of ampicillin at 100 mg/mL and kanamycin at 50 mg/mL can be premade, filter-sterilized, and stored in aliquots at -20 °C. Dilute these 100x in 1x PBS to obtain the working solution.
4. Using a pipette, remove as much liquid as possible from the centrifuge tube, leaving the larva behind, and add 90 µL of the 1x PBS with antibiotics.
   NOTE: Antibiotics are used to prevent bacterial growth in GMM plates that may interfere with the counting of Aspergillus colonies.
5. Homogenize larvae in a tissue lyser at 1,800 oscillations/min (30 Hz) for 6 min. Spin down at 17,000 x g for 30 s.
6. Label GMM plates (one plate per homogenized larvae). Using a Bunsen burner to create a sterile environment, pipette the homogenized suspension from one tube to the middle of the GMM plate, then spread using a disposable L-shaped spreader. Avoid spreading the homogenate against the rim.
7. Incubate the plates upside down at 37 °C for 2–3 days and count the number of colonies formed, colony forming unit (CFU).
8. To measure the number of spores alive during the infection period, pick larvae from the 96-well plates at 1–7 days post injection (dpi) and transfer them to centrifuge tubes. Euthanize and homogenize larvae to spread on GMM plates as described in steps 4.1–4.5.

5. Drug treatment of injected larvae

1. After section 4, split the remaining injected larvae into two 3.5 mm dishes: one for the drug treatment and one for the control. Use about 24 infected larvae per condition.
   NOTE: The 3.5 mm dishes can be treated with 2% nonfat dry milk in water, rinsed, air-dried, and stored at RT beforehand. Coating with milk will prevent larvae from sticking to the plastic.
2. Prepare the desired drug solution and the vehicle in E3 without methylene blue in conical tubes according to the final concentration required, then mix well. For example, to monitor the survival of larvae exposed to dexamethasone, use 24 larvae (replicates) for dexamethasone and 24 for vehicle control, such as dimethyl sulfoxide (DMSO). Prepare 5 mL of the drug solution at the required concentration. Here, 5 mL of 0.1% DMSO and 10 µM dexamethasone were used, and 24 larvae/condition were transferred to ~200 µL of the vehicle/drug solution/larvae.
3. Remove as much liquid as possible from one dish with a transfer pipette and add premixed E3 containing vehicle control. Repeat with premixed E3 containing the treatment of interest for the other dish.
4. Using a pipette, transfer larvae into a 96-well plate (one larva per well). Monitor the survival of injected larvae exposed to the vehicle or the drug for 7 days.
   NOTE: The drug can be applied solely on the day of infection and kept on the larvae for the entire experiment or can be refreshed daily.

6. Daily imaging of infected larvae using the zebrafish Wounding and Entrapment Device for Growth and Imaging (zWEDGI)

1. Ensure that larvae are treated with 100 µM N-phenylthiourea (PTU) at 24 hpf to prevent pigmentation and that PTU is kept on the larvae for the entire experiment.
   NOTE: PTU at 75–100 µM prevents pigmentation of larvae without any gross developmental defects. However, PTU can interfere with some biological processes, and researchers should determine beforehand whether the drug may affect any processes under investigation.
2. Infect transgenic larvae with labeled cell populations of interest at 2 dpf with Aspergillus spores engineered to express a fluorescent protein, as described in section 3. Then, transfer infected larvae into wells of a 48-well plate in about 500 µL/well of E3 without methylene blue.
   NOTE: A 48-well plate is used here because it is easier to transfer larvae into and out of during repeated daily imaging.
3. On the day of imaging, prepare two 3.5 mm Petri dishes: one with 100 µM PTU and one with E3-tricaine.
4. Add E3-tricaine into the chambers of a zWEDGI device. Under the stereo microscope, remove air bubbles from the chambers and the restraining channel using a P100 micropipette. Remove all excess E3-tricaine, so that is it only in the chambers.
5. Pipette up one larva from the plate using a transfer pipette. If a lot of liquid is used to remove it, pipette it into a 3.5 mm dish containing E3-PTU. Then, pipette up again, using as little liquid as possible, and transfer into E3-tricaine.
6. Wait for ~30 s for anesthetization, then transfer into the loading chamber of the wounding and entrapment device (e.g., zWEDGI).
7. Under the stereo microscope, position the larva. Use the P100 micropipette to remove E3-tricaine from the wounding chamber and release it into the loading chamber to move the tail of the larva into the restriction channel. Ensure that the larva is positioned on its lateral, dorsal, or dorsolateral side so that the hindbrain can be imaged with an inverted objective lens.
8. Image larva with a confocal microscope.
9. After imaging, with the P100 pipette, release E3-tricaine into the wounding chamber to push the larva from the restraining channel into the loading chamber.
10. Using a transfer pipette, pick up the larva and transfer it back to the Petri dish with E3-Tricaine. Using as little liquid as possible, transfer it to the Petri dish with E3-PTU. Rinse in PTU and transfer back into the 48-well plate.