Using Zebrafish Models of Human Influenza A Virus Infections to Screen Antiviral Drugs and Characterize Host Immune Cell Responses

The overall goals of these procedures are to use zebrafish as an influenza model to study the host immune response to viral infection and to screen antiviral compounds. These methods can help answer key questions in the influenza field, such as the role phagocytes play in immunity and inflammation during an infection and the efficacy of potential antiviral therapies. A zebrafish model of influenza allows for previously unanswered questions to be addressed due to the optical clarity of developing zebrafish and the capacity to perform medium to high through-put assays.

After preparing reagants and raising embryos to the desired stage according to the text protocol, on the day of the experiment, use number five forceps to dechorionate the embryos. If using the APR-8 or X31 viruses, in a laminar flow hood, dilute the viruses to three point three times 10 to the six egg infective dose 50 percent in sterile ice cold PBS supplemented with zero point two five percent phenol red. If using the NS1-GFP strain, dilute the virus to approximately one point five times 10 to the second PFUs per nanoliter.

Simultaneously, set up a control containing allantoic fluid from uninfected chicken eggs diluted similar to the virus. Then, with microloader tips, pipette virus solution into microinjection needles and insert the microinjection needle into the appropriate holder of the injection apparatus. After clipping the needle according to the text protocol, adjust the pressure and timing settings on the pressure injector and or reclip the needle tip until the desired injection volume is achieved.

Following anesthetization of embryos, use a plastic pipette to transfer 10 to 20 embryos to a two percent agarose plate and pipette to remove excess liquid. Under the stereomicroscope, use a fire polished and sealed borosilicate glass capillary to gently align and orient the embryos so that the duct of cuvier or the posterior cardinal vein is in line with the microinjection needle. To carry out microinjection of IAV, gently insert the microinjection needle into the duct of cuvier or the posterior cardinal vein.

Then, depress the foot pedal to inject the desired volume of IAV into the circulatory system of the embryo. To achieve a systemic infection, it is critical that the injection bolus is swept up into the circulation and distributed throughout the body. If the bolus collects at the site of infection, remove the embryo from the experiment and euthanize it.

Five days post fertilization, prepare needles as described earlier in this video. Align TG MPX mCherry larvae containing inflated swim bladders as just demonstrated, except position the larvae so that the needle can pierce the swim bladder and the virus can be deposited into the posterior of the swim bladder. Gently insert the microinjection needle into the swim bladder and inject the desired volume towards the posterior of the organ.

The injection solution should collect towards the posterior and the swim bladder’s air bubble should be displaced forward. To achieve a swim bladder infection, it is critical that the injection bolus collects towards the posterior of the swim bladder, causing the air bubble to be displaced forward. GFP expression should begin to be observed as early as three hours post injection.

After completing the injections, including with control solutions, transfer the larvae to Petri dishes containing sterile egg water and incubate the fish at 33 degrees Celsius. To carry out antiviral drug treatment, at three hours post infection, replace the egg water of the NS1-GFP and control infected fish with sterile water containing zero, 16.7, or 33.3 nanograms per milliliter of Zanamivir. Beginning at 24 to 48 hours post injection, begin to record observations about infection dynamics related to disease pathology, including signs of lethargy and evidence of edema, differences in pigmentation, ocular and cranial facial deformities, and lordosis.

Every 24 hours for five days post infection, record the number of dead, morbid, and healthy larvae. Plot data as a stacked bar graph with percentages of healthy, morbid, or dead fish plotted on the Y axis and the treatment group on the x axis. To mount zebrafish for imaging, transfer an individual transgenic MPX mCherry larvae that has been infected with NS1-GFP to a well of a 24 well glass bottom plate and a small droplet of egg water.

Repeat with other IAV infected and control fish. Slowly add one percent agarose embryo mounting medium to each well, taking care not to introduce large bubbles. Gently adjust the position of the larva so that each one is mounted on its side.

Once the agarose has gelled, use egg water with 200 micrograms per liter of tricaine to gently fill the individual wells. With a confocal microscope and a 20 times objective focused on the swim bladder, capture a z stack series of brightfield and fluorescence images. Quantify and compare the number of MPX mCherry positive neutrophils present in the swim bladders of IAV infected and control injected zebrafish.

Shown here are data representing how systemic IAV infection in zebrafish can be used to test drug efficacy. Embryos at 48 HPF were injected with APR8, X31 or NS1-GFP via the duct of cuvier to initiate a viral infection. Embryos serving as controls for viral infection were injected at 48 HPF.

By 48 hours post infection, zebrafish injected with IAV exhibited evidence of pericardial edema and circulatory arrest with erythrocytes present throughout the pericardium. Infected fish not exposed to Zanamivir exhibited features of edema in systemic GFP expression. Reduced gross pathology in GFP florescence were observed in NS1-GFP infected larvae exposed to Zanamivir.

In this experiment of a localized zebrafish swim bladder infection model, NS1-GFP virus was injected into the swim bladders of five day post fertilization transgenic MPX mCherry larvae and the recruitment of mCherry labeled neutrophils into the swim bladder was tracked. At 20 hours post infection, considerable migration of neutrophils into the swim bladders of IAV infected fish relative to PBS injected controls was observed, indicating that IAV can recruit neutrophils to the swim bladder, just as it recruits neutrophils to the human lung. The implications of these techniques extend towards antiviral therapies because they have the potential to reveal critical insights into host influenza A virus or IAV interactions that may ultimately translate into the clinic.

Don’t forget that working with influenza can be extremely hazardous, and precautions including vaccination and wearing the appropriate PPE should always be taken while performing this procedure.