This work was supported by the National Institute of Allergy And Infectious Diseases of the National Institutes of Health under Award Number K22AI134677. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Researchers should obtain approval for all animal experiments from the appropriate animal care and use committees. Representative data shown in this article are from experiments performed under protocols approved by the Clemson University Institutional Animal Care and Use Committee (AUP2018-070, AUP2019-012).
1. Preparation of Aspergillus spores for injection
<ol>
	<li>From an Aspergillus spore suspension, calculate the volume needed to obtain 1 x 106 spores. The volume should be 20&#8211;100 &#956;L; 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 &#181;L, produce a 10x dilution and use 50 &#181;L of the diluted solution. 
	NOTE: Two plates/strain can be prepared to collect more spores or as a spare in the case of contamination.</li>
	<li>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 &#176;C for 3&#8211;4 days, with the plate facing upside down.</li>
	<li>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.</li>
	<li>Place one piece of miracloth in each labeled 50 mL conical tube and re-cap. Take the remaining miracloth packet out of the hood.</li>
	<li>Bring plates into the biosafety cabinet. Open one plate, then pour Tween-water on the top to cover about three-quarters of the plate.</li>
	<li>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 &#8220;puffs&#8221; 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 extraction of different strains.</li>
	<li>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.</li>
	<li>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&#160;and pour them into the same conical tube.</li>
	<li>Pour Tween-water to bring the total volume in the conical tube to 50 mL.</li>
	<li>Spin at 900 x g for 10 min. Make sure to use aerosolization-preventing caps in the centrifuge.</li>
	<li>Pour off the supernatant into ~10% bleach solution to decontaminate. Pour 50 mL of sterile 1x PBS into the conical tube, then vortex or shake to resuspend the pellet.</li>
	<li>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.</li>
	<li>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 &#181;L of the fungal homogenate with 900 &#181;L of Tween-water).</li>
	<li>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.</li>
	<li>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</li>
	<li>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 &#176;C for ~4 weeks.</li>
	<li>Prior to use in injections, mix 20 &#181;L of the spore preparation with 10 &#181;L 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.</li>
	<li>For a mock injection, mix 20 &#181;L of 1x PBS with 10 &#181;L of 1% sterile phenol red.</li>
</ol>
2. Preparation of agar plates for injection
<ol>
	<li>Prepare 2% agarose in E3 medium and melt in a microwave.</li>
	<li>Pour into a 100 mm x 15 mm Petri dish (~25 mL per plate), swirl to cover the plate evenly, and let cool.</li>
	<li>Wrap the plate with paraffin film and store inverted at 4 &#176;C.</li>
	<li>Prior to injection, bring the plate to room temperature (RT).</li>
	<li>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 &#176;C. 2% BSA pre-treatment prevents larvae from sticking to the surface of the agarose.</li>
	<li>Pour E3 with buffered tricaine onto the plate and let it sit until injection.</li>
</ol>
3. Zebrafish larva hindbrain ventricle microinjection
<ol>
	<li>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.</li>
	<li>Remove as much E3 as possible from the Petri dish and add buffered 300 &#181;g/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 &#730;C. The working solution can be made by diluting 4 mL of the stock solution up to 50 mL with E3.</li>
	<li>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).</li>
	<li>Open the compressed air valve and turn on the microinjector. Set the pressure to ~25 PSI, pulse duration to 60 ms, and backpressure unit to 1&#160;PSI.</li>
	<li>Load a microinjection needle using a microloader pipette tip (Table of Materials) with about 3&#8211;5 &#181;L of prepared PBS or spore suspension with phenol red. Mount the needle onto the micromanipulator. 
	NOTE: Microinjection needles can be prepared as described previously23. The stereo microscope used for microinjections should have an eye piece reticle to calibrate the microinjection needle. The reticle should be calibrated with a stage micrometer, and the length of the reticle scale (&#181;m) 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.</li>
	<li>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.</li>
	<li>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).</li>
	<li>Move micromanipulator and needle out of the way to avoid accidentally hitting the needle while the larvae are arranged on the injection plate.</li>
	<li>Pour E3-Tricaine off the injection plate, then transfer ~24 anesthetized larvae to the injection plate with as little E3 possible using a transfer pipet.</li>
	<li>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 &#8220;float&#8221; 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.</li>
	<li>Adjust the microscope zoom to the lowest magnification. Bring the micromanipulator back and arrange so that the needle is close to the larvae, at a ~30&#730;&#8211;60&#730; angle, in the middle of the field of view.</li>
	<li>Zoom in to the highest magnification and use fine adjustment knobs to further adjust the position of the needle. Verify that ~30&#8211;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 after every five to six larvae, as the number of spores coming out of the needle can increase or decrease over time.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Moving down the plate, inject all the larvae in that row. Then, turn the plate around and inject all larvae in 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.</li>
	<li>Move 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.</li>
	<li>First, pull away with 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.</li>
	<li>Repeat as necessary for the final experimental sample number desired.</li>
	<li>Rinse larvae at least 2x with E3 and ensure recovery from anesthesia.</li>
	<li>To quantify survival without any further treatment, using a transfer pipette, transfer larvae into a 96 well plate (1 larva per well) in E3.</li>
</ol>
4. Establishment of injected and viable spore numbers
<ol>
	<li>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).</li>
	<li>Euthanize larvae with tricaine or by placing them at 4 &#176;C for 0.5&#8211;2.0 h.</li>
	<li>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 &#176;C and used later. 
	NOTE: Stock solutions of ampicillin at 100 mg/mL and kanamycin 50 mg/mL can be premade, filter-sterilized, and stored in aliquots at -20 &#176;C. Dilute these 100x in 1x PBS to obtain the working solution.</li>
	<li>Using a pipette, remove as much liquid as possible from the centrifuge tube, leaving the larva behind and add 90 &#181;L of the 1x PBS with antibiotics. 
	NOTE: Antibiotics are used to prevent bacterial growth in GMM plates that may interfere with counting of Aspergillus colonies.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Incubate the plates upside down at 37 &#176;C for 2&#8211;3 days and count the number of colonies formed (CFU).</li>
	<li>To measure the number of spores alive during the infection period, pick larvae from the 96 well plate at 1&#8211;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&#8211;4.5.</li>
</ol>
5. Drug treatment of injected larvae
<ol>
	<li>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 sticking to the plastic.</li>
	<li>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 the vehicle control, such as DMSO. Prepare 5 mL of the drug solution at the required concentration. Here, 5 mL of 0.1% DMSO and 10 &#181;M dexamethasone were used, and 24 larvae/condition were transferred to ~200 &#181;L of the vehicle/drug solution/larvae.</li>
	<li>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.</li>
	<li>Using a pipette, transfer larvae into 96 well plate (one larva per well). Monitor the survival of injected larvae exposed to the vehicle or the drug for 7 days. 
	NOTE: Drug can be applied solely on the day of infection and kept on the larvae for the entire experiment or can be refreshed daily.</li>
</ol>
6. Daily imaging of infected larvae using the zebrafish Wounding and Entrapment Device for Growth and Imaging (zWEDGI)
<ol>
	<li>Ensure that larvae are treated with 100 &#181;M 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&#8211;100 &#181;M prevents pigmentation of larvae without any gross developmental defects24. However, PTU can interfere with some biological processes25, and researchers should determine beforehand whether the drug may affect any processes under investigation.</li>
	<li>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 &#181;L/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.</li>
	<li>On the day of imaging, prepare two 3.5 mm Petri dishes: one with 100 &#181;M PTU and one with E3-tricaine.</li>
	<li>Add E3-tricaine into the chambers of a zWEDGI device26,27. 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.</li>
	<li>Pipette up one larva from the plate using a transfer pipette. If a lot of liquid is used to remove it, pipette into a 3.5 mm dish containing E3-PTU. Then, pipette up again, using as little liquid as possible, and transfer into E3-tricaine.</li>
	<li>Wait ~30 s for anesthetization, then transfer into the loading chamber of the wounding and entrapment device (e.g., zWEDGI).</li>
	<li>Under the stereo microscope, position the larva. Use the P100 micropipette to remove E3-tricaine from the wounding chamber and release into the loading chamber to move the tail of the larva into the restriction channel. Ensure that larva is positioned on its lateral, dorsal, or dorso-lateral side, so that the hindbrain can be imaged with an inverted objective lens.</li>
	<li>Image larva with a confocal microscope.</li>
	<li>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.</li>
	<li>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.</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenylthiourea+disrupts+thyroid+function+in+developing+zebrafish.">Phenylthiourea disrupts thyroid function in developing zebrafish.</a> Development Genes and Evolution. 212 (12), 593-598 (2003).</li><li>Huemer, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=zWEDGI:+Wounding+and+Entrapment+Device+for+Imaging+Live+Zebrafish+Larvae.">zWEDGI: Wounding and Entrapment Device for Imaging Live Zebrafish Larvae.</a> Zebrafish. 14 (1), 42-50 (2017).</li><li>Huemer, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+Live+Imaging+Device+for+Improved+Experimental+Manipulation+of+Zebrafish+Larvae.">Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae.</a> Journal of Visualized Experiments. (128), e56340 (2017).</li><li>Perlin, D. S., Shor, E., Zhao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Update+on+Antifungal+Drug+Resistance.">Update on Antifungal Drug Resistance.</a> Current Clinical Microbiology Reports. 2 (2), 84-95 (2015).</li></ol>

No conflicts or financial interests to disclose.

The infection model described here is beneficial for analyzing the host immune responses, host-pathogen interactions, and fungal pathogenesis12,13,14,15. This information can be derived from the high-resolution imaging of fluorescent-labeled pathogens and host cells13, larval survival, and CFU persistence over time.
The microinjection technique is critical to the success of this protocol and may need to be adjusted when using different microinjection equipment and setups. In particular, the pressure and time of injection are two major variables and can be adjusted to ensure that the volume ejected by the needle is ~3 nL. The size of the needle as determined by clipping it with forceps also regulates the number of spores being injected; although, a larger opening can cause tissue damage to the larva. On the other hand, too small of an opening will not allow the relatively large spores (&#62;2 &#181;m) out and can lead to needle clogging. If this occurs, the needle can be reclipped to have a slightly larger opening.
Other protocols for microinjection of bacteria utilize PVP-40 to help maintain a homogenous injection mixture, but we have not found any advantage in using this carrier with Aspergillus spores. Clogging of the needle can be mitigated by vortexing the fungal preparation thoroughly to break any clumps prior to loading the needle. Sometimes, a clog in the needle can also be dislodged by temporarily increasing the pressure or injection time and triggering the microinjector while the needle is in the liquid surrounding the larvae. The pressure and injection time should then be decreased again to previous levels. In other cases, a clog cannot be removed, and a new needle needs to be loaded and recalibrated.
This protocol is designed to inject ~30&#8211;70 spores per larva. It is known that based on the concentration of the spore preparation and the volume injected, this number is quite low. However, it has been empirically found that this is the number of spores injected under these conditions. Why this difference occurs is unknown, but it may be due to spore clumping in the needle. Our own attempts to inject larger numbers of spores have largely been unsuccessful.
To ensure about 30&#8211;70 spores are being injected and maintain the consistency of the injections throughout all the larvae, check the number of spores by injecting onto the E3 surrounding the larvae. Repeat this every five to six larvae throughout all the injections. If the spore count seems to change, the pressure and/or injection time can be adjusted to inject a consistent number of spores across multiple larvae. However, care should be taken that the injection dose remains primarily in the hindbrain and does not fill the midbrain and forebrain.
To ensure a localized infection, the spore suspension should be contained within the hindbrain ventricle. This can be visualized by the phenol red staining just after the injection, though the red color diffuses with time. For injections, the region around the otic vesicle is used to pierce through and reach the ventricle at a 45&#176;&#8211;65&#176; angle. This area has no main blood vessels, causes less tissue damage, and heals instantly. If the skin over the ventricle is pierced, the spore suspension can be leaked out, because the needle that must be used for Aspergillus spore injections is larger than that is used for bacterial suspensions. Unsuccessfully injected or accidentally damaged larvae can be marked by injecting into the yolk a couple of times to create a red mark or by dragging the larva out of the row with the needle. After a set of injections is complete, these larvae should be removed and disposed of before the rest are washed off the plate. E3 without methylene blue is used to anesthetize larvae prior to injection and also keep larva after the injections, because methylene blue is anti-fungal.
At the time of injection, CFU counts represent the number of viable spores within the infected host. However, if the spores germinate into hyphae, these can be broken up into separate viable &#8220;fungal units&#8221; during homogenization and can give rise to multiple colonies. Or, an unbroken multicellular hypha can give rise to a single colony, resulting in an averaged, but imprecise, representation of the fungal burden. This can be mitigated by combining the CFU counts with longitudinal microscopy of individual larvae, which provides visual data of the fate of injected spores.
Compared to the mammalian system, the zebrafish larva infection model is particularly significant due to its optical accessibility. The recruitment and response of innate immune cells can be visualized within a live intact host. This can be incorporated with genetic or chemical inhibition of molecular targets to analyze how each target affects the macrophage or neutrophil reaction against Aspergillus spores in a live animal.
While the zebrafish larva Aspergillus infection model continues to be instrumental in describing different aspects of IA12,13,14,15,16,17,18,19,20,22, there are other areas of expansion. From the host side, it is used to describe cellular level immune responses, but this can be expanded to analyze immune mechanisms at the molecular level by combining it with targeted morpholino, CRISPR, stable mutant lines, or chemical exposure. One caveat is that homologues for all known mammalian innate immune pathway components have not been identified in zebrafish.
From the pathogen side, virulence of different species and strains have been described. A promising avenue of future research is the use of mutant Aspergillus strains to test how specific genes or proteins contribute as virulence factors. Thereby, novel anti-fungal drugs can be developed to target these proteins. Current anti-fungal drugs have low efficacy in human patients and there is growing resistance to these drugs in fungi28. This in vivo model can be used to investigate why these drugs fail and as an intermediate model to test the efficacy of novel anti-fungal drugs. Overall, the findings discovered using this model can facilitate future development of effective treatments for Aspergillus-infected patients.

Aspergillus fumigatus is a ubiquitous saprophytic fungus, and its airborne spores can be found both indoors and outdoors1. These spores are inhaled by everyone but become effectively cleared from the lungs of immunocompetent individuals1,2. However, people with altered lung conditions such as cystic fibrosis can develop bronchopulmonary aspergillosis due to fungal germination in the lungs3. The most severe form of this infection, invasive aspergillosis (IA), affects immunocompromised individuals and involves growth of the fungus into other organs2,3. IA leads to &#62;50% death of infected patients despite the availability of anti-fungal therapies4. In immunocompetent individuals, innate immune responses play a major role in clearing the inhaled spores1. However, the specific mechanisms that contribute to this innate immune clearance are not well-understood. It is important to understand the cellular and molecular mechanisms of major innate immune cells (i.e., macrophages and neutrophils) in clearance of Aspergillus in order to find novel therapeutic strategies for IA.
While mammalian models have been instrumental in identifying fungal virulence factors and host immune responses5,6, visual accessibility is limited for host-pathogen interactions at the cellular level. Tissue culture experiments cannot fully recapitulate the complex multi-cellular environment and interactions that exist in whole animals7. Therefore, zebrafish has gained popularity as an alternative model organism to fill this gap and facilitate the study of host-pathogen interactions in a live, intact host across a multi-day infection8,9. The zebrafish innate immune system develops as early as 24 h post-fertilization (hpf)10, and the adaptive system takes 4&#8211;6 weeks to develop11, providing a window of time in which innate immune responses can be assessed in isolation. Innate immune responses are well-conserved between humans and zebrafish11. Zebrafish have many qualities that facilitate the investigation of these responses, including optical clarity (which allows for the high-resolution live imaging of intact hosts) and genetic tractability (which facilitates molecular mechanistic studies).
The larval zebrafish Aspergillus infection model described here was originally developed by Knox et al.12. It has recently been expanded by our group and others to investigate host immune mechanisms12,13, host-pathogen interactions13,14,15, mechanisms of immunosuppression13,16,17, fungal virulence18, and anti-fungal drug efficacy19,20. This model recapitulates multiple aspects of human aspergillosis. While immunocompetent larvae are resistant, immunocompromised larvae can succumb to infection12,13,16,17.
In this model, a localized infection is established by injecting spores into the hindbrain ventricle of larva, an area less populated with phagocytes, and phagocyte recruitment and behavior can be evaluated12,13. It is believed that macrophages act as the first line of defense against Aspergillus spores in humans1 and mammalian models6,21. Similarly, in the zebrafish model, macrophages are recruited to the injected Aspergillus spores, while neutrophils are recruited secondarily in response to hyphal growth12,13,22. From this model, it has also been learned that Aspergillus can persist in wildtype immunocompetent larvae after more than 7 days of infection. Furthermore, the entire course of the infection can be followed in the same live animals by daily confocal imaging.
This protocol describes the technique of microinjection to inject spores into the hindbrain ventricle of 2 days post-fertilization (2 dpf) larvae. The infection is then monitored for up to 7 days, as zebrafish larvae can live up to 10 dpf without feeding. Immunosuppression can be induced by drug treatment, and the application of drugs to the larvae is also described. Finally, two methods to follow infection progression are described, including quantification of CFUs from individual larvae and a daily live imaging setup.

<table>
	<tbody>
		<tr>
			<td>Dumont forceps #5</td>
			<td>Roboz Surgical Instrument Co.</td>
			<td>RS-5045</td>
			<td></td>
		</tr>
		<tr>
			<td>Eyepiece reticle</td>
			<td>Microscope World</td>
			<td>RETR10</td>
			<td>For calibrating needles, used in Stereomicroscope</td>
		</tr>
		<tr>
			<td>Microinjector setup: Back pressure unit</td>
			<td>Applied Scientific Instrumentation</td>
			<td>BPU</td>
			<td></td>
		</tr>
		<tr>
			<td>Footswitch</td>
			<td>Applied Scientific Instrumentation</td>
			<td>FTSW</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro pipet holder kit</td>
			<td>Applied Scientific Instrumentation</td>
			<td>M-Pip</td>
			<td></td>
		</tr>
		<tr>
			<td>Pressure injector</td>
			<td>Applied Scientific Instrumentation</td>
			<td>MPPI-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator setup: Micromanipulator</td>
			<td>Narashige (Tritech)</td>
			<td>M-152</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetic stand and plate</td>
			<td>Tritech</td>
			<td>MINJ-HBMB</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle puller</td>
			<td>Sutter Instrument</td>
			<td>P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Nikon</td>
			<td>SMZ-745</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissuelyser II</td>
			<td>Qiagen</td>
			<td>85300</td>
			<td>To homogenize larvae</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments/Description</td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Fisher</td>
			<td>BP160-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin sodium salt</td>
			<td>Fisher</td>
			<td>AAJ6380706</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA, fraction V</td>
			<td>VWR</td>
			<td>AAJ65855-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Kanamycin sulfate</td>
			<td>Fisher</td>
			<td>AAJ1792406</td>
			<td></td>
		</tr>
		<tr>
			<td>L spreaders</td>
			<td>Fisher</td>
			<td>14 665 230</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcapillary needles (no filament)</td>
			<td>World Precision Instruments (WPI)</td>
			<td>TW100-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Microloader pipet tips</td>
			<td>VWR</td>
			<td>89009-310</td>
			<td>To load the needle with Aspergillus suspension</td>
		</tr>
		<tr>
			<td>Miracloth</td>
			<td>VWR</td>
			<td>EM475855-1R</td>
			<td>To filter Aspergillus suspension</td>
		</tr>
		<tr>
			<td>N-phenylthiourea</td>
			<td>Fisher</td>
			<td>AAL0669009</td>
			<td>To prevent pigmentation</td>
		</tr>
		<tr>
			<td>Phenol red, 1% solution</td>
			<td>Fisher</td>
			<td>57254</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine (Ethyl 3-aminobenzoate, methanesulfonic acid salt)</td>
			<td>Fisher</td>
			<td>AC118000500</td>
			<td>To anesthetize larvae</td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Fisher</td>
			<td>BP337-500</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Media and Solutions</td>
			<td>Components/Recipe</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>E3 media: 60x E3</td>
			<td>17.2 g NaCl, 0.76 g KCl, 2.9 g CaCl2, 4.9 g MgSO4 &#183; 7H2O, to 1 L with H2O</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1x E3</td>
			<td>16.7 ml 60x stock, 430 ul 0.05 M NaOH, to 1 L with H2O (optional: + 3 ml 0.01% methylene blue)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine stock solution</td>
			<td>2 g Tricaine, 5 g Na2HPO4 &#183; 7H2O, 4.2 ml 60X E3, to 500 ml with H2O, pH to 7.0-7.5 with NaOH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose minimal media (GMM) agar: GMM agar</td>
			<td>10 g Glucose (Dextrose), 50 ml 20x Nitrate salts, 1 ml TE, to 1 L with H2O, pH to 6.5 with NaOH, + 16 g Agar, autoclave</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>20x Nitrate salts</td>
			<td>120 g NaNO3, 10.4 g KCl, 10.4 g, MgSO4 &#183; 7H2O, 30.4 g, KH2PO4, to 1 L with H2O, autoclave</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trace elements (TE)</td>
			<td>2.20 g ZnSO4 &#183; 7H2O, 1.10 g H3BO3, 0.50 g MnCl2 &#183; 4H2O, 0.16 g FeSO4 &#183; 7H2O, 0.16 g CoCl2 &#183; 6H2O, 0.16 g CuSO4 &#183; 5H2O, 0.11 g (NH4)6Mo7O24 &#183; 4H2O, 5.00 g Na2EDTA, to 100 ml with H2O, dissolve stirring overnight, autoclave</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

infection of zebrafish larvae with aspergillus spores for analysis of host-pathogen interactions

Invasive aspergillosis (IA) is one of&#160;the most common fungal infections among immunocompromised individuals. Despite the availability of antifungal drugs, IA can cause &#62;50% mortality in infected immunocompromised patients. It is crucial to determine both host and pathogen factors that contribute to infection susceptibility and low survival rates in infected patients in order to develop novel therapeutics. Innate immune responses play a pivotal role in recognition and clearance of Aspergillus spores, though little is known about the exact cellular and molecular mechanisms. Reliable models are required to investigate detailed mechanistic interactions between the host and pathogen. The optical clarity and genetic tractability of zebrafish larvae make them an intriguing model&#160;to study host-pathogen interactions of multiple human bacterial and fungal infections in a live and intact host. This protocol describes a larval zebrafish Aspergillus infection model. First, Aspergillus spores are isolated and injected into the zebrafish hindbrain ventricle via microinjection. Then, chemical inhibitors such as immunosuppressive drugs are added directly to the larval water. Two methods to monitor the infection in injected larvae are described, including the 1) homogenization of larvae for colony forming unit (CFU) enumeration and 2) a repeated, daily live imaging setup. Overall, these techniques can be used to mechanistically analyze the progression of Aspergillus infection in vivo and can be applied to different host backgrounds and Aspergillus strains to interrogate host-pathogen interactions.

After microinjection of Aspergillus spores into the hindbrain of zebrafish larvae, infection outcome can be followed by multiple assays, including survival, CFUs, and live imaging. In a survival assay, the number of infected larvae surviving 1&#8211;7 dpi was monitored. When wildtype larvae were left untreated, very little death was observed, with ~80%&#8211;100% of larvae surviving the entirety of the experiment (Figure 1). If larvae were immunosuppressed, such as by exposure to the corticosteroid drug dexamethasone (10 &#181;M), significantly decreased survival was observed (Figure 1).
When CFUs were quantified throughout the 7 day experiment from wildtype larvae infected with A. fumigatus spores, persistence of spores was observed, with slow clearance over time (Figure 2A). The number of spores surviving at 1, 2, 3, 5, and 7 dpi were normalized to the number of spores injected at 0 dpi to compare persistence and clearance across replicates (Figure 2B).
Transgenic fish lines expressing fluorescent proteins in leukocytes together with fluorescent protein-expressing Aspergillus spores can be used to visualize both leukocyte recruitment and behavior as well as fungal germination and growth13. When macrophages were labeled (e.g., Tg(mpeg1:H2B-GFP)), macrophage clustering in ~50% of larvae was typically observed, starting at 2&#8211;3 dpi (Figure 3A). Neutrophil (Tg(lyz:BFP)) recruitment was typically delayed, occurring primarily after fungal germination has occurred (Figure 3A). While fungal burden persisted for the whole experiment in the majority of larvae (Figure 3A), clearance was observed (Figure 3B). In some larvae, fungal burden outside of the hindbrain was also observed later in infection, due to fungal dissemination, likely in macrophages.
The area around the otic vesicle is one possible location where this dissemination can&#160;be&#160;found (Figure 3C). These observations were quantified in multiple individual larvae over the course of the entire experiment (Figure 4). Typically, germination was seen in ~60% of larvae by 5 dpi (Figure 4A). Phagocyte cluster area, macrophage recruitment, and neutrophil recruitment vary both over time and across larvae, with some trending up throughout the experiment and some resolving over time (Figure 4B,C,D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61165/61165fig01.jpg" /> 
Figure 1: Representative survival analysis of infected larvae.&#160;Aspergillus-infected larvae were exposed to vehicle control (DMSO) or dexamethasone (Dex), and survival was monitored. Data represent three pooled replicates. Average injection CFUs: DMSO = 30, Dex = 29 (p-value and hazard ratio were calculated by Cox proportional hazard regression analysis, ****p &#60; 0.0001). <a href="https://www.jove.com/files/ftp_upload/61165/61165fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61165/61165fig2v2.jpg" /> 
Figure 2: Representative CFU counts from individual infected larvae immediately after injection (0 dpi) and during infection course (2, 3, 5, and 7 dpi).&#160;Eight infected larvae were homogenized and plated to count CFU for each timepoint and replicate. (A) Example data from one replicate. Each point represents one larva, bars represent means for each timepoint. (B) CFU counts were normalized to the CFU count at 0 dpi for each replicate, and three replicates were pooled. Data were compared between experimental conditions using analysis of variance and summarized in terms of estimated marginal means and standard errors. Asterix represent statistical significance compared to CFU at 0 dpi (*p &#60; 0.05, **p &#60; 0.01, ****p &#60; 0.0001). <a href="https://www.jove.com/files/ftp_upload/61165/61165fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61165/61165fig3v2.jpg" /> 
Figure 3: Representative images from infection experiments.&#160;PTU-treated larvae with fluorescent macrophages (mpeg1:H2B-GFP) and neutrophils (lyz:BFP) were injected with RFP-expressing A. fumigatus. Live, infected larvae were imaged repeatedly at 2, 3, and 5 dpi on a confocal microscope. Maximum intensity Z projection images are displayed. Schematics of larvae indicate location of imaging for each panel. All scale bars represent 100 &#181;m, except for the inset scale bars, which are 25 &#181;m. (A) Images shown are from a single larva taken at 2, 3, and 5 dpi, representing a typical infection progression. Insets show fungal germination at days 3 and 5. (B) Representative image of subset of larvae that can clear the infection, with low fungal burden and not much inflammation at 5 dpi. (C) Representative image of subset of larvae in which infection disseminates out of the hindbrain at later timepoints. In this image, fungus, macrophages, and neutrophils can be found around and below the otic vesicle. <a href="https://www.jove.com/files/ftp_upload/61165/61165fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61165/61165fig4v2.jpg" /> 
Figure 4: Representative quantification of imaging experiments.&#160;Images from experimental setup in Figure 3 were analyzed for fungal germination and leukocyte recruitment. (A) Larvae were scored for the presence of germinated spores on each day, and the percentage of larvae with germination was calculated. (B,C,D) Each individual larva is represented as a different color line. Phagocyte cluster appearance and size (B), macrophage recruitment (C), and neutrophil recruitment (D) were followed over the course of the 5 day experiment for each larva. <a href="https://www.jove.com/files/ftp_upload/61165/61165fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Infection of Zebrafish Larvae with Aspergillus Spores for Analysis of Host-Pathogen Interactions at JoVE.com

Infection of Zebrafish Larvae with Aspergillus Spores for Analysis of Host-Pathogen Interactions

This protocol describes an Aspergillus infection model in zebrafish larvae. Aspergillus spores are microinjected into the hindbrain of larvae, and chemical treatment is used to induce immunosuppression. Infection progression is monitored via a daily imaging setup to monitor fungal growth and immune responses as well as enumeration of live spores by colony forming unit plating.

Infection of Zebrafish Larvae with Aspergillus Spores for Analysis of Host-Pathogen Interactions

Invasive aspergillosis (IA) is one of&#160;the most common fungal infections among immunocompromised individuals. Despite the availability of antifungal ...

infection-zebrafish-larvae-with-aspergillus-spores-for-analysis-host

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Immunology and Infection

4.0K Views.  Clemson University. This protocol describes an Aspergillus infection model in zebrafish larvae. Aspergillus spores are microinjected into the hindbrain of larvae, and chemical treatment is used to induce immunosuppression. Infection progression is monitored via a daily imaging setup to monitor fungal growth and immune responses as well as enumeration of live spores by colony forming unit plating.

Video: Infection of Zebrafish Larvae with Aspergillus Spores for Analysis of Host-Pathogen Interactions

Use of an Optical Trap for Study of Host-Pathogen Interactions for Dynamic Live Cell Imaging

Dynamic live cell imaging allows direct visualization of real-time interactions between cells of the immune system1, 2; however, the lack of spatial and temporal control between the phagocytic cell and microbe has rendered focused observations into the initial interactions of host response to pathogens difficult. Historically, intercellular contact events such as phagocytosis3 have been imaged by mixing two cell types, and then continuously scanning the field-of-view to find serendipitous intercellular contacts at the appropriate stage of interaction. The stochastic nature of these events renders this process tedious, and it is difficult to observe early or fleeting events in cell-cell contact by this approach. This method requires finding cell pairs that are on the verge of contact, and observing them until they consummate their contact, or do not. To address these limitations, we use optical trapping as a non-invasive, non-destructive, but fast and effective method to position cells in culture.
Optical traps, or optical tweezers, are increasingly utilized in biological research to capture and physically manipulate cells and other micron-sized particles in three dimensions4. Radiation pressure was first observed and applied to optical tweezer systems in 19705, 6, and was first used to control biological specimens in 19877. Since then, optical tweezers have matured into a technology to probe a variety of biological phenomena8-13.
We describe a method14 that advances live cell imaging by integrating an optical trap with spinning disk confocal microscopy with temperature and humidity control to provide exquisite spatial and temporal control of pathogenic organisms in a physiological environment to facilitate interactions with host cells, as determined by the operator. Live, pathogenic organisms like Candida albicans and Aspergillus fumigatus, which can cause potentially lethal, invasive infections in immunocompromised individuals15, 16 (e.g. AIDS, chemotherapy, and organ transplantation patients), were optically trapped using non-destructive laser intensities and moved adjacent to macrophages, which can phagocytose the pathogen. High resolution, transmitted light and fluorescence-based movies established the ability to observe early events of phagocytosis in living cells. To demonstrate the broad applicability in immunology, primary T-cells were also trapped and manipulated to form synapses with anti-CD3 coated microspheres in vivo, and time-lapse imaging of synapse formation was also obtained. By providing a method to exert fine spatial control of live pathogens with respect to immune cells, cellular interactions can be captured by fluorescence microscopy with minimal perturbation to cells and can yield powerful insight into early responses of innate and adaptive immunity.

A method is described to individually select, manipulate, and image live pathogens using an optical trap coupled to a spinning disk microscope. The optical trap provides spatial and temporal control of organisms and places them adjacent to host cells. Fluorescence microscopy captures dynamic intercellular interactions with minimal perturbation to cells.

Dynamic live cell imaging allows direct visualization of real-time interactions between cells of the immune system1, 2; however, the lack of spatial and ...

Use of the EpiAirway Model for Characterizing Long-term Host-pathogen Interactions

Nontypeable Haemophilus influenzae (NTHi) are human-adapted Gram-negative bacteria that can cause recurrent and chronic infections of the respiratory mucosa 1; 2. To study the mechanisms by which these organisms survive on and inside respiratory tissues, a model in which successful long-term co-culture of bacteria and human cells can be performed is required. We use primary human respiratory epithelial tissues raised to the air-liquid interface, the EpiAirway model (MatTek, Ashland, MA). These are non-immortalized, well-differentiated, 3-dimensional tissues that contain tight junctions, ciliated and nonciliated cells, goblet cells that produce mucin, and retain the ability to produce cytokines in response to infection. 
This biologically relevant in vitro model of the human upper airway can be used in a number of ways; the overall goal of this method is to perform long-term co-culture of EpiAirway tissues with NTHi and quantitate cell-associated and internalized bacteria over time. As well, mucin production and the cytokine profile of the infected co-cultures can be determined. This approach improves upon existing methods in that many current protocols use submerged monolayer or Transwell cultures of human cells, which are not capable of supporting bacterial infections over extended periods3. For example, if an organism can replicate in the overlying media, this can result in unacceptable levels of cytotoxicity and loss of host cells, arresting the experiment. The EpiAirway model allows characterization of long-term host-pathogen interactions. Further, since the source for the EpiAirway is normal human tracheo-bronchial cells rather than an immortalized line, each is an excellent representation of actual human upper respiratory tract tissue, both in structure and in function4.
For this method, the EpiAirway tissues are weaned off of anti-microbial and anti-fungal compounds for 2 days prior to delivery, and all procedures are performed under antibiotic-free conditions. This necessitates special considerations, since both bacteria and primary human tissues are used in the same biosafety cabinet, and are co-cultured for extended periods.

This method allows characterization of extended bacterial co-culture with EpiAirways, primary human respiratory epithelial tissue grown at the air-liquid interface, a biologically relevant in vitro model. The approach can be used with any microbe that is amenable to long-term co-culture.

Nontypeable Haemophilus influenzae  (NTHi) are human-adapted Gram-negative bacteria that can cause recurrent and chronic infections of the respiratory ...

Infection of Zebrafish Embryos with Intracellular Bacterial Pathogens

Zebrafish (Danio rerio) embryos are increasingly used as a model for studying the function of the vertebrate innate immune system in host-pathogen interactions 1. The major cell types of the innate immune system, macrophages and neutrophils, develop during the first days of embryogenesis prior to the maturation of lymphocytes that are required for adaptive immune responses. The ease of obtaining large numbers of embryos, their accessibility due to external development, the optical transparency of embryonic and larval stages, a wide range of genetic tools, extensive mutant resources and collections of transgenic reporter lines, all add to the versatility of the zebrafish model. Salmonella enterica serovar Typhimurium (S. typhimurium) and Mycobacterium marinum can reside intracellularly in macrophages and are frequently used to study host-pathogen interactions in zebrafish embryos. The infection processes of these two bacterial pathogens are interesting to compare because S. typhimurium infection is acute and lethal within one day, whereas M. marinum infection is chronic and can be imaged up to the larval stage 2, 3. The site of micro-injection of bacteria into the embryo (Figure 1) determines whether the infection will rapidly become systemic or will initially remain localized. A rapid systemic infection can be established by micro-injecting bacteria directly into the blood circulation via the caudal vein at the posterior blood island or via the Duct of Cuvier, a wide circulation channel on the yolk sac connecting the heart to the trunk vasculature. At 1 dpf, when embryos at this stage have phagocytically active macrophages but neutrophils have not yet matured, injecting into the blood island is preferred. For injections at 2-3 dpf, when embryos also have developed functional (myeloperoxidase-producing) neutrophils, the Duct of Cuvier is preferred as the injection site. To study directed migration of myeloid cells towards local infections, bacteria can be injected into the tail muscle, otic vesicle, or hindbrain ventricle 4-6. In addition, the notochord, a structure that appears to be normally inaccessible to myeloid cells, is highly susceptible to local infection 7. A useful alternative for high-throughput applications is the injection of bacteria into the yolk of embryos within the first hours after fertilization 8. Combining fluorescent bacteria and transgenic zebrafish lines with fluorescent macrophages or neutrophils creates ideal circumstances for multi-color imaging of host-pathogen interactions. This video article will describe detailed protocols for intravenous and local infection of zebrafish embryos with S. typhimurium or M. marinum bacteria and for subsequent fluorescence imaging of the interaction with cells of the innate immune system.

Transparent zebrafish embryos have proved useful model hosts to visualize and functionally study interactions between innate immune cells and intracellular bacterial pathogens, such as Salmonella typhimurium and Mycobacterium marinum. Micro-injection of bacteria and multi-color fluorescence imaging are essential techniques involved in the application of zebrafish embryo infection models.

Zebrafish (Danio rerio) embryos are increasingly used as a model for studying the function of the vertebrate innate immune system in host-pathogen ...

Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 to 40% attributable mortality6. Systemic candidiasis is normally controlled by innate immunity, and individuals with genetic defects in innate immune cell components such as phagocyte NADPH oxidase are more susceptible to candidemia7-9. Very little is known about the dynamics of C. albicans interaction with innate immune cells in vivo. Extensive in vitro studies have established that outside of the host C. albicans germinates inside of macrophages, and is quickly destroyed by neutrophils10-14. In vitro studies, though useful, cannot recapitulate the complex in vivo environment, which includes time-dependent dynamics of cytokine levels, extracellular matrix attachments, and intercellular contacts10, 15-18. To probe the contribution of these factors in host-pathogen interaction, it is critical to find a model organism to visualize these aspects of infection non-invasively in a live intact host. 
The zebrafish larva offers a unique and versatile vertebrate host for the study of infection. For the first 30 days of development zebrafish larvae have only innate immune defenses2, 19-21, simplifying the study of diseases such as disseminated candidiasis that are highly dependent on innate immunity. The small size and transparency of zebrafish larvae enable imaging of infection dynamics at the cellular level for both host and pathogen. Transgenic larvae with fluorescing innate immune cells can be used to identify specific cells types involved in infection22-24. Modified anti-sense oligonucleotides (Morpholinos) can be used to knock down various immune components such as phagocyte NADPH oxidase and study the changes in response to fungal infection5. In addition to the ethical and practical advantages of using a small lower vertebrate, the zebrafish larvae offers the unique possibility to image the pitched battle between pathogen and host both intravitally and in color. 
The zebrafish has been used to model infection for a number of human pathogenic bacteria, and has been instrumental in major advances in our understanding of mycobacterial infection3, 25. However, only recently have much larger pathogens such as fungi been used to infect larva5, 23, 26, and to date there has not been a detailed visual description of the infection methodology. Here we present our techniques for hindbrain ventricle microinjection of prim25 zebrafish, including our modifications to previous protocols. Our findings using the larval zebrafish model for fungal infection diverge from in vitro studies and reinforce the need to examine the host-pathogen interaction in the complex environment of the host rather than the simplified system of the Petri dish5.

The rapid development, small size and transparency of zebrafish are tremendous advantages for the study of innate immune control of infection1-4. Here we demonstrate techniques for infecting zebrafish larvae using the fungal pathogen Candida albicans by microinjection, methodology recently used to implicate phagocyte NADPH oxidase activity in control of fungal dimorphism5.

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 ...

A Comparative Approach to Characterize the Landscape of Host-Pathogen Protein-Protein Interactions

Significant efforts were gathered to generate large-scale comprehensive protein-protein interaction network maps. This is instrumental to understand the pathogen-host relationships and was essentially performed by genetic screenings in yeast two-hybrid systems. The recent improvement of protein-protein interaction detection by a Gaussia luciferase-based fragment complementation assay now offers the opportunity to develop integrative comparative interactomic approaches necessary to rigorously compare interaction profiles of proteins from different pathogen strain variants against a common set of cellular factors.
This paper specifically focuses on the utility of combining two orthogonal methods to generate protein-protein interaction datasets: yeast two-hybrid (Y2H) and a new assay, high-throughput Gaussia princeps protein complementation assay (HT-GPCA) performed in mammalian cells.
A large-scale identification of cellular partners of a pathogen protein is performed by mating-based yeast two-hybrid screenings of cDNA libraries using multiple pathogen strain variants. A subset of interacting partners selected on a high-confidence statistical scoring is further validated in mammalian cells for pair-wise interactions with the whole set of pathogen variants proteins using HT-GPCA. This combination of two complementary methods improves the robustness of the interaction dataset, and allows the performance of a stringent comparative interaction analysis. Such comparative interactomics constitute a reliable and powerful strategy to decipher any pathogen-host interplays.

This article focuses on the identification of high-confident interaction datasets between host and pathogen proteins using a combination of two orthogonal methods: yeast two-hybrid followed by a high-throughput interaction assay in mammalian cells called HT-GPCA.

Significant efforts were gathered to generate  large-scale comprehensive protein-protein interaction network maps. This is  instrumental to understand the ...

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte extravasation might result in pathophysiological changes in the CNS. Thus, investigation of lymphocyte migration into the CNS is important to understand inflammatory CNS diseases and to develop new therapy approaches. Here we present an in vitro model of the human blood-brain barrier to study lymphocyte extravasation. Human brain microvascular endothelial cells (HBMEC) are confluently grown on a porous polyethylene terephthalate transwell insert to mimic the endothelium of the blood-brain barrier. Barrier function is validated by zonula occludens immunohistochemistry, transendothelial electrical resistance (TEER) measurements as well as analysis of evans blue permeation. This model allows investigation of the diapedesis of rare lymphocyte subsets such as CD56brightCD16dim/- NK cells. Furthermore, the effects of other cells, cytokines and chemokines, disease-related alterations, and distinct treatment regimens on the migratory capacity of lymphocytes can be studied. Finally, the impact of inflammatory stimuli as well as different treatment regimens on the endothelial barrier can be analyzed.

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Here, we describe a human blood-brain barrier model enabling to investigate lymphocyte transmigration into the central nervous system in vitro.

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte ...

Ex Vivo Infection of Human Lymphoid Tissue and Female Genital Mucosa with Human Immunodeficiency Virus 1 and Histoculture

Histocultures allow studying intercellular interactions within human tissues, and they can be employed to model host-pathogen interactions under controlled laboratory conditions. Ex vivo infection of human tissues with human immunodeficiency virus (HIV), among other viruses, has been successfully used to investigate early disease pathogenesis, as well as a platform to test the efficacy and toxicity of antiviral drugs. In the present protocol, we explain how to process and infect with HIV-1 tissue explants from human tonsils and cervical mucosae, and maintain them in culture on top of gelatin sponges at the liquid-air interface for about two weeks. This non-polarized culture setting maximizes access to nutrients in culture medium and oxygen, although progressive loss of tissue integrity and functional architectures remains its main limitation. This method allows monitoring HIV-1 replication and pathogenesis using several techniques, including immunoassays, qPCR, and flow cytometry. Of importance, the physiologic variability between tissue donors, as well as between explants from different areas of the same specimen, may significantly affect experimental results. To ensure result reproducibility, it is critical to use an adequate number of explants, technical replicates, and donor-matched control conditions to normalize the results of the experimental treatments when compiling data from multiple experiments (i.e., conducted using tissue from different donors) for statistical analysis.

Ex Vivo Infection of Human Lymphoid Tissue and Female Genital Mucosa with Human Immunodeficiency Virus 1 and Histoculture

Infection of human tissues with human immunodeficiency virus (HIV) ex vivo provides a valuable 3D model of virus pathogenesis. Here, we describe a protocol to process and infect tissue specimens from human tonsils and female genital mucosae with HIV-1 and maintain them in culture at the liquid-air interface.

Histocultures allow studying intercellular interactions within human tissues, and they can be employed to model host-pathogen interactions under ...

Extraction of Hemocytes from Drosophila melanogaster Larvae for Microbial Infection and Analysis

During the pathogenic infection of Drosophila melanogaster, hemocytes play an important role in the immune response throughout the infection. Thus, the goal of this protocol is to develop a method to visualize the pathogen invasion in a specific immune compartment of flies, namely hemocytes. Using the method presented here, up to 3 &#215; 106 live hemocytes can be obtained from 200 Drosophila 3rd instar larvae in 30 min for ex vivo infection. Alternatively, hemocytes can be infected in vivo through injection of 3rd instar larvae followed by hemocyte extraction up to 24 h post-infection. These infected primary cells were fixed, stained, and imaged using confocal microscopy. Then, 3D representations were generated from the images to definitively show pathogen invasion. Additionally, high-quality RNA for qRT-PCR can be obtained for the detection of pathogen mRNA following infection, and sufficient protein can be extracted from these cells for Western blot analysis. Taken together, we present a method for definite reconciliation of pathogen invasion and confirmation of infection using bacterial and viral pathogen types and an efficient method for hemocyte extraction to obtain enough live hemocytes from Drosophila larvae for ex vivo and in vivo infection experiments.

Extraction of Hemocytes from Drosophila melanogaster Larvae for Microbial Infection and Analysis

This method demonstrates how to visualize pathogen invasion into insect cells with three-dimensional (3D) models. Hemocytes from Drosophila larvae were infected with viral or bacterial pathogens, either ex vivo or in vivo. Infected hemocytes were then fixed and stained for imaging with a confocal microscope and subsequent 3D cellular reconstruction.

During the pathogenic infection of Drosophila melanogaster, hemocytes play an important role in the immune response throughout the infection. Thus, the ...

Evaluation of Host-Pathogen Responses and Vaccine Efficacy in Mice

Vaccines are a 20th century medical marvel. They have dramatically reduced the morbidity and mortality caused by infectious diseases and contributed to a striking increase in life expectancy around the globe. Nonetheless, determining vaccine efficacy remains a challenge. Emerging evidence suggests that the current acellular vaccine (aPV) for Bordetella pertussis (B. pertussis) induces suboptimal immunity. Therefore, a major challenge is designing a next-generation vaccine that induces protective immunity without the adverse side effects of a whole-cell vaccine (wPV). Here we describe a protocol that we used to test the efficacy of a promising, novel adjuvant that skews immune responses to a protective Th1/Th17 phenotype and promotes a better clearance of a B. pertussis challenge from the murine respiratory tract. This article describes the protocol for mouse immunization, bacterial inoculation, tissue harvesting, and analysis of immune responses. Using this method, within our model, we have successfully elucidated crucial mechanisms elicited by a promising, next-generation acellular pertussis vaccine. This method can be applied to any infectious disease model in order to determine vaccine efficacy.

Here we present an elegant protocol for in vivo evaluation of vaccine effectiveness and host immune responses. This protocol can be adapted for vaccine models that study viral, bacterial, or parasitic pathogens.

Vaccines are a 20th century medical marvel. They have dramatically reduced the morbidity and mortality caused by infectious diseases and contributed to a ...

Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry

Invasive pulmonary infection by the mold Aspergillus fumigatus poses a great threat to immunocompromised patients. Inhaled fungal conidia (spores) are cleared from the human lung alveoli by being phagocytosed by innate monocytes and/or neutrophils. This protocol offers a fast and reliable measurement of phagocytosis by flow cytometry using fluorescein isothiocyanate (FITC)-labeled conidia for co-incubation with human leukocytes and subsequent counterstaining with an anti-FITC antibody to allow discrimination of internalized and cell-adherent conidia. Major advantages of this protocol are its rapidness, the possibility to combine the assay with cytometric analysis of other cell markers of interest, the simultaneous analysis of monocytes and neutrophils from a single sample and its applicability to other cell wall-bearing fungi or bacteria. Determination of percentages of phagocytosing leukocytes provides a means to microbiologists for evaluating virulence of a pathogen or for comparing pathogen wildtypes and mutants as well as to immunologists for investigating human leukocyte capabilities to combat pathogens.

Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry

This protocol provides a fast and reliable method to quantitatively measure phagocytosis of Aspergillus fumigatus conidia by human primary phagocytes using flow cytometry and to discriminate phagocytosis of conidia from mere adhesion to leukocytes.

Invasive pulmonary infection by the mold Aspergillus fumigatus poses a great threat to immunocompromised patients. Inhaled fungal conidia (spores) are ...