The authors would like to thank the laboratory of Dr. Carol Kim for microinjection training, Clarissa Henry for advice on speeding up embryo development and use of equipment, and Nathan Lawson for contributing fli1:EGFP fish. We thank members of the Wheeler lab and Shawn Walls for critical reading of the manuscript. We would also like to thank Mark Nilan for fish care and advice, and Ryan Phennicie and Kristin Gabor for technical advice on this project. This work was funded by a MAFES research assistantship to K. Brothers, a MAFES Hatch grant E08913-08, and a NIH NCRR award P20RR016463 to R. Wheeler.

All zebrafish care protocols and experiments were performed under Institutional Animal Care and Use Committee (IACUC) protocol A2009-11-01.
1. Morpholino and Larval Injection Dishes
Experimental duration: * (10-15 minutes)
Degree of difficulty: *
<ol>
	<li>For egg injections, prepare a 2% agarose solution in sterile water and microwave. When the solution has cooled pour some of it into an extra deep Petri dish (Fisher Scientific) until it is half full. Cool on ice and make sure the plate is level.</li>
	<li>Once the dish has cooled, pour a 15 mL top layer of 2% agarose. Spray an egg injection grooved plastic mold (Adaptive Science Tools) with sterile water from a spray bottle. Carefully place the mold groove side down into the hot agarose. When the agarose has cooled, use a metal flat spatula (VWR Scientific) to dissociate the mold from the agarose. Slowly remove the grid from the agarose. Wrap embryo injection dishes in Parafilm (VWR Scientific) and store inverted at 4 &deg;C.</li>
	<li>For fish larval injections, prepare a 2% agarose solution as described. Pour the solution into a standard size Petri dish (VWR Scientific) and set aside until it solidifies. Wrap larval injection dishes in Parafilm and store inverted at 4 &deg;C.</li>
</ol>
2. Fungal Culture Preparation
Experimental duration: **(30 minutes)
Degree of difficulty: **
<ol>
	<li>Prepare yeast extract-peptone-dextrose (YPD) agar plates: 10 g/liter yeast extract, 20 g/liter peptone, 20 g/liter dextrose, and 20 g/liter agar into and autoclave. For YPD liquid prepare 10 g/liter yeast extract, 20 g/liter peptone, 20 g/liter dextrose and autoclave 20-30 minutes at 121 &deg;C.</li>
	<li>Two days prior to fish infections prepare a streak plate from frozen stocks of Candida albicans cultures onto YPD agar to obtain single colonies.</li>
	<li>Incubate at 37 &deg;C overnight.</li>
	<li>Put 5 mL YPD broth into 16 x 150 mm culture tubes (VWR Scientific).</li>
	<li>The day before the fish infections pick 1 small colony on the YPD agar with a wooden dowel (VWR Scientific). Put the stick in the culture tube and swirl around to re-suspend colony in the YPD liquid.</li>
	<li>Grow overnight at 37 &deg;C on a TC-7 Tissue Culture Roller Drum equipped with a 14-inch test tube wheel (New Brunswick Scientific).</li>
	<li>The next day, spin down 1 mL of culture at 14000x g for 1 minute in a 1.7 mL tube (Axygen). Remove the supernatant and resuspend the pellet in residual liquid by vortexing (VWR Scientific).</li>
	<li>Add 1 mL 1x Phosphate Buffered Saline (VWR Scientific) and spin as previously described.</li>
	<li>Repeat the PBS wash 3 times.</li>
	<li>Dilute 1:100 dilution in 1x PBS (10 &mu;L of washed C. albicans into 990 &mu;L 1x PBS).</li>
	<li>Count on a Hemocytometer (VWR Scientific).</li>
	<li>Dilute to 107 cells/mL.</li>
</ol>
3. Zebrafish Infections
Experimental duration: **** (1-3 hours)
Degree of difficulty: ****
<ol>
	<li>Collect embryos according to section 5 and store in egg-water, 60 mg/L instant ocean salts (Fisher Scientific) in sterile deionized water.</li>
	<li>Using a dissecting microscope such as the Olympus SZ61 (Olympus), dechorionate embryos on infection day. Use Dumont Dumoxel tweezers (VWR Scientific) to pull the chorion27 apart like opening a bag of chips, or gently poke the tweezers in the closed position into the chorion and then slowly open them. The fish should pop right out of the chorion.</li>
	<li>Swirl the extra deep Petri dish with the lid on to move fish into the center of the dish. Transfer fish to the lid of the dish. Remove egg-water and replace with fresh media. Add fish to media.</li>
	<li>Warm larval injection dishes in an incubator at 28 &deg;C. Prepare tricaine methane sulfonate (Western Chemical Inc.) dilution at 200 &mu;g/mL for anesthetizing fish.</li>
	<li>Count out the desired number of larvae for infection (20-50 fish). Put fish in tricaine methane sulfonate solution and wait 1-2 minutes until they stop moving.</li>
	<li>Turn on the MPPI-3 injection unit (Applied Scientific Instruments). Make sure the pressure switch is on "pulse" and the "pulse duration" is set to 9 with 3 PSI for the backpressure unit. Open the valve to the nitrogen tank until the pressure on the injection unit reads 30 PSI.</li>
	<li>Load a pulled micropipette28 with 5 &mu;L of vortexed Candida albicans at a concentration of 1x107 cells/mL. Place the micropipette in the micropipette holder (Applied Scientific Instruments). Fill an empty extra deep Petri dish with water. Move the micropipette until the tip is within view just touching the surface of the water via dissecting microscope and use Dumont Dumoxel tweezers (VWR Scientific) to clip the needle about 3mm from the tip of the pulled pipette.</li>
	<li>Press the foot switch (Applied Scientific Instruments) to verify the needle has been clipped. You should see the liquid disperse if you were successful. The diameter of the liquid bolus should be no bigger than the diameter of the pupil of the prim25 zebrafish larva (0.21 mm, yielding a sphere of 4.9 nL in volume). Adjust the pressure accordingly if the liquid bolus is too big or too small.</li>
	<li>Anesthetize the fish in tricaine methane sulfonate (200 &mu;g/mL). Once the fish have stopped moving, swirl the dish with the lid on until the fish are in the center. Collect 50 fish using a transfer pipette (Fisher Scientific). Tap the side of the pipette to settle the fish towards the tip. Gently pipette fish onto the larval agarose dish using as little liquid as possible.</li>
	<li>Line the fish up with a smooth glass rod (be careful not to crush them!). Aspirate as much liquid as possible off of the dish. Use a KimWipe to wick away any moisture.</li>
	<li>Position the larval agarose dish with fish under the microscope until you get a good view of the fish and the needle at the same time. Move the glass needle towards the fish. Zoom in on both the fish and needle as you position the needle towards the fish. Carefully move the glass needle into the otic vesicle27, 29 (ear) of the first fish. You'll know once you've moved the needle into the fish because when you push the foot switch (Applied Scientific Instruments), the hindbrain ventricle will lift up slightly.</li>
	<li>Press the foot switch (Applied Scientific Instruments) and watch the liquid disperse inside the fish's hindbrain ventricle27. Retract the needle from the fish and move to the next fish. Repeat procedure until you have injected all of the fish on the plate. Remove any dead fish and inject replacements to keep the number correct (50 fish). For each group of 50 fish injected a new microinjection needle must be prepared, or the C. albicans settles in the bottom of the needle and clogs it, or throws off the concentration injected.</li>
	<li>Wash the fish off of the dish by angling the dish and spraying egg-water onto the fish into a clean extra deep Petri dish containing 60 mL egg-water. It is important not to leave the fish on the agarose for longer than 15-20 minutes or they will dry out and die.</li>
	<li>Repeat entire injection process until finished. Don't forget PBS and non-pathogenic injected controls.</li>
	<li>When finished, keep the fish at 28 &deg;C.</li>
	<li>Close the nitrogen tank valve. Move the injection unit switch from "pulse" to "continuous" to relieve pressure in the tank line, the pressure should drop to zero at this point. Switch injection unit back to "pulse". Turn off the injection unit and clean up the work area.</li>
</ol>
4. Preparing the Fish for Imaging
Experimental duration: ** (30 minutes)
Degree of difficulty: **
<ol>
	<li>Prepare tricaine methane sulfonate solution as before (200 &mu;g/mL). Get out infected fish, and transfer them into tricaine.</li>
	<li>Prepare 47.9 mL 0.4% low-melt agarose (VWR scientific) in egg-water. Heat the solution by microwaving. Cool to 37 &deg;C and add tricaine methane sulfonate (200 &mu;g/mL) to mixture</li>
	<li>Once fish are immobilized in tricaine methane sulfonate, pipette individual fish into a Petri dish (VWR scientific) containing 0.4% low melt agarose (VWR Scientific).</li>
	<li>Next, move the fish from low melt agarose into individual wells of a glass bottom dish (MatTek Corporation) for imaging. Use as little low melt agarose as possible so the fish lies flat on the bottom of the dish. Only use enough of the tricaine-agarose to fill the inner circles of the glass bottom dish.</li>
</ol>
5. Modifications Related to JoVE Protocols
Micropipettes for Microinjection
Experimental duration: * (10-15 minutes)
Degree of difficulty: *
<ol>
	<li>Pull hollow glass rods BF120-69-10 (Sutter Instruments) using a Flaming Brown Micropipette Puller Model P-97 (Sutter Instruments) according to Yuan et al.30. Choose program #7 with the following conditions Heat = 470, Velocity = 120, Time = 200. The resulting needle is 8 mm from bevel to tip and, once clipped at 3 mm above the tip it has a diameter of approximately 10 &mu;m.</li>
	<li>Load a glass micropipette into the brackets. Select "Pull." The heating filament will heat the needle according to the program parameters and the glass rod will separate into two pulled micropipettes. Store micropipettes in a pipette storage box (Sutter instruments).</li>
</ol>
Embryo Collection, Morpholino Injection and Maintenance
Experimental duration: *** (1-2 hours)
Degree of difficulty: ***
<ol start="3">
	<li>Collect embryos according to the methods of Rosen et al.31. Use a plastic sieve (Wares of Knutsford) to collect eggs from the spawning tank (Aquatic Habitats). Hold the sieve upside down over an extra deep Petri dish and rinse with tank water to collect eggs.</li>
	<li>For morpholino preparation, add 300 &mu;L sterile water to 300 nanomoles of morpholino (GeneTools, LLC). This gives a working stock of 1.0 mM.</li>
	<li>Verify the concentration of the morpholino by using a Nanodrop (Thermo Scientific). Select "nucleic acid and change the sample type to "other" and wavelength to 265. Enter the constant by multiplying the molecular weight of the morpholino by 1000 divided by the absorptivity coefficient listed in the morpholino oligo property sheet.</li>
	<li>Blank the nanodrop with 2 &mu;L 0.1 N HCl. Dilute 5 &mu;L of the morpholino solution into 95 &mu;L of 0.1 N HCl (20x dilution). Place 2 &mu;L of the dilution on the nanodrop pedestal and click measure. Multiply the concentration by the dilution factor (20). This gives a working concentration of morpholino. To calculate the millimolar concentration divide the concentration obtained by the molecular weight of the morpholino.</li>
	<li>Prepare a working stock of morpholino in Danieau buffer (58 mM NaCl, 0.7 mM KCl, 0.4 mM MgSO4, 0.6 mM Ca(NO3)2, 5.0 mM HEPES, pH 7.6) and 0.01% phenol red (VWR Scientific).</li>
	<li>For morpholino injection work follow the protocol according to Rosen et al.31 and Yuan et al.30. Rinse morpholino injection dishes with egg-water and warm at 28 &deg;C for 15 minutes. Remove water from injection dishes and line-up one-cell stage embryos into the grooves of the injection dish (prepared from Section 1) with a transfer pipette. Each plate will hold approximately 250 eggs. Inject 1-2 cell stage embryos with morpholino. Eggs will remain in the one-cell stage for 15 minutes27.</li>
	<li>Rinse injected eggs with egg-water when finished (sterile deionized water with 60 mg/L Instant Ocean salts) and place in 60 mL egg-water in extra deep Petri dishes. Morpholino injection dishes can be reused. Rinse with egg-water when finished and store inverted at 4 &deg;C.</li>
	<li>For storage of embryos add 60 mL egg-water to dish. Count out 110 embryos with a transfer pipette into separate extra deep Petri dishes with 60 mL egg-water. Add 0.00003% methylene blue to prevent microbial growth and incubate embryos at 28 &deg;C.</li>
	<li>To speed up development of embryos keep embryo dishes at 33 &deg;C for 24 hours and stage according to Kimmel et al.27, allowing embryos to develop to a head trunk angle of 75&deg; (Prim 25 stage27).</li>
	<li>For larval infection work keep embryos at 28 &deg;C after injecting with C. albicans.</li>
	<li>Each day replace embryo dishes with 60 mL fresh egg-water.</li>
</ol>
Imaging
Experimental duration: *****(1-5 hours)
Degree of difficulty: ***
<ol start="14">
	<li>Prepare fish in low melt agarose (according to section 4.2) with tricaine in glass bottom imaging dishes (MatTek Corporation) as described earlier. Place the dish on an Olympus IX-81 (Olympus) inverted microscope stage. Bring the fish into focus under 4x magnification under differential interference contrast (DIC) light. Images can be captured at 4x, 20x, and 40x magnification in DIC, TRITC (tetramethyl rhodamine isothiocyanate), and FITC (fluorescein isothiocyanate) filter settings.</li>
	<li>Use an Olympus IX-81 inverted microscope with an FV-1000 laser scanning confocal system according to Ariga et al.32. Conduct time courses at 40x magnification by setting Z-stacks with 1-1.5 &mu;m slices every hour of infection.</li>
	<li>For long time course imaging of 2 hours or more, place a layer of low melt agarose on top of the fish in the imaging dish every 2-3 hours. This prevents the agarose from drying out and crushing the fish while it is being imaged. For lengthy time courses with a small number of fish, use a heated stage (Bioptechs Inc.) to maintain 28 &deg;C during infections.</li>
</ol>
6. Representative Results
An example of a successful hindbrain ventricle C. albicans infection in a zebrafish larva at 5 hours post-infection (hpi) and 24 hpi is shown in (Figure 1). Macrophage-like cells with engulfed C. albicans are seen in the hindbrain ventricle at 5 hpi. By 24 hpi, C. albicans is inside macrophage-like cells in the dorsal tail tissue indicative of disseminated candidiasis. This infection result is highly dependent upon an accurate injection of 10-15 yeast-form C. albicans into the hindbrain ventricle. Screening of infected fish immediately post-injection can ensure this.
<img alt="Figure 1" src="/files/ftp_upload/4051/4051fig1.jpg" /> 
Figure 1. Transgenic fli1:EGFP22, 33 larva infected with CAF2-yCherry Candida albicans and imaged intravitally by confocal microscopy. (A-C) 5 hours post-infection (A) Infected larva with EGFP-expressing macrophage-like cells at the site of infection (hindbrain ventricle) Scale bar =100 &mu;m. (B and C) Higher magnification images of same fish, showing C. albicans within phagocytes. Scale bar = 100 &mu;m for B and 10 &mu;m for C. (D-F) 24 hours post-infection (D) Infected larva with disseminated candidiasis with CAF2-yCherry C. albicans inside EGFP macrophage-like cells in the dorsal tail tissue. Scale bar = 100 &mu;m. (E and F) Higher magnification images of same fish, showing C. albicans in tail tissue. Scale bar = 100 &mu;m.

No conflicts of interest declared.

The zebrafish microinjection method presented here differs from Gutzman et al.34 in that here we demonstrate injection through the otic vesicle into the hindbrain ventricle of 36 to 48 hpf larvae. The method we describe allows for consistent injection of 10-15 yeast into the hindbrain ventricle with reduced tissue damage. This protocol produces an initially local infection that spreads throughout the body by 24 hpi (Figure 1) and results in significant lethality/morbidity5</su...

<table border="1">
<tbody>
 <tr>
 <td>Name of the 			reagent</td>
 <td>Company</td>
 <td>Catalog 			number</td>
 <td>Comments 			(optional)</td>
 </tr>
 <tr>
 <td>Spawning tanks</td>
 <td>Aquatic 			habitats</td>
 <td>&nbsp;2L</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>1.7 mL tubes</td>
 <td>Axygen</td>
 <td>MCT-175-C</td>
 <td></td>
 </tr>
 <tr>
 <td>Instant Ocean</td>
 <td>Fisher 			Scientific</td>
 <td>S17957C </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Extra deep 			Petri dishes</td>
 <td>Fisher 			Scientific</td>
 <td>08-757-11Z</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Standard Petri 			dishes</td>
 <td>VWR Scientific</td>
 <td>25384-302</td>
 <td></td>
 </tr>
 <tr>
 <td>Transfer 			pipettes</td>
 <td>Fisher 			Scientific</td>
 <td>13-711-7M</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Yeast Extract</td>
 <td>VWR Scientific</td>
 <td>90000-726</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Peptone</td>
 <td>VWR Scientific</td>
 <td>90000-264</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dextrose</td>
 <td>Fisher 			Scientific</td>
 <td>D16-1</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Agar</td>
 <td>VWR Scientific</td>
 <td>90000-760</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Disposable 			Hemocytometer</td>
 <td>VWR Scientific</td>
 <td>82030-468</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Phosphate 			Buffered Saline</td>
 <td>VWR Scientific</td>
 <td>12001-986</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dumont Dumoxel 			Tweezers</td>
 <td>VWR Scientific</td>
 <td>100501-806</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Wooden Dowels</td>
 <td>VWR Scientific</td>
 <td>10805-018</td>
 <td></td>
 </tr>
 <tr>
 <td>KimWipes</td>
 <td>VWR Scientific</td>
 <td>300053-964</td>
 <td></td>
 </tr>
 <tr>
 <td>Low Melt 			Agarose</td>
 <td>VWR Scientific</td>
 <td>12001-722</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Agarose for 			injection dishes</td>
 <td>VWR Scientific</td>
 <td>12002-102</td>
 <td></td>
 </tr>
 <tr>
 <td>Flaming Brown 			Micropipette Puller</td>
 <td>Sutter 			Instruments</td>
 <td>P-97</td>
 <td></td>
 </tr>
 <tr>
 <td>Hollow glass 			rods</td>
 <td>Sutter 			Instruments</td>
 <td>BF120-69-10</td>
 <td>For glass rods 			smooth glass by heating over bunsen burner&nbsp;</td>
 </tr>
 <tr>
 <td>Pipette 			Storage Box</td>
 <td>Sutter 			Instruments</td>
 <td>BX10</td>
 <td></td>
 </tr>
 <tr>
 <td>MPPI-3 			Injection system</td>
 <td>Applied 			Scientific Instrumentation</td>
 <td>MPPI-3</td>
 <td></td>
 </tr>
 <tr>
 <td>Back Pressure 			Unit</td>
 <td>Applied 			Scientific Instrumentation</td>
 <td>BPU</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micropipette 			Holder kit</td>
 <td>Applied 			Scientific Instrumentation</td>
 <td>MPIP</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Foot Switch</td>
 <td>Applied 			Scientific Instrumentation</td>
 <td>FSW</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micromanipulator</td>
 <td>Applied 			Scientific Instrumentation</td>
 <td>MM33</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Magnetic Base</td>
 <td>Applied 			Scientific Instrumentation</td>
 <td>Magnetic Base</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tricaine 			methane sulfonate</td>
 <td>Western 			Chemical Inc.</td>
 <td>MS-222</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dissecting 			Scope</td>
 <td>Olympus</td>
 <td>SZ61 top 			SZX-ILLB2-100 base</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Confocal 			Microscope</td>
 <td>Olympus</td>
 <td>IX-81 with 			FV-1000 laser scanning confocal system</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>TC-7 Tissue 			Culture Roller 			drum with 14 inch test tube wheel</td>
 <td>New Brunswick 			Scientific</td>
 <td>&nbsp;TC-7</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Imaging Dishes</td>
 <td>MatTek 			Corporation</td>
 <td>P24G-1.0-10-F</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Pipette tips for 			loading needles</td>
 <td>Eppendorf</td>
 <td>930001007</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Plate pouring 			grids</td>
 <td>Adaptive 			Science Tools</td>
 <td>TU-1</td>
 <td></td>
 </tr>
 <tr>
 <td>Heated Stage</td>
 <td>Bioptechs Inc.</td>
 <td>Delta T-5</td>
 <td></td>
 </tr>
 <tr>
 <td>Flat Spatula</td>
 <td>VWR Scientific</td>
 <td>82027-486</td>
 <td></td>
 </tr>
 <tr>
 <td>Plastic Sieves</td>
 <td>Wares of 			Knutsford Online</td>
 <td>12 cm</td>
 <td></td>
 </tr>
 <tr>
 <td>Parafilm</td>
 <td>VWR Scientific</td>
 <td>52858-000 </td>
 <td></td>
 </tr>
 <tr>
 <td>Vortex Genie</td>
 <td>VWR Scientific</td>
 <td>14216-184</td>
 <td></td>
 </tr>
 <tr>
 <td>16 x 150 mm 			Culture tubes</td>
 <td>VWR Scientific</td>
 <td>60825-435</td>
 <td></td>
 </tr>
 <tr>
 <td>Nanodrop</td>
 <td>Thermo 			Scientific</td>
 <td>ND 2000</td>
 <td></td>
 </tr>
 <tr>
 <td>Phenol Red</td>
 <td>VWR Scientific</td>
 <td>97062-478 </td>
 <td></td>
 </tr>
 <tr>
 <td>HCl</td>
 <td>VWR Scientific</td>
 <td>87003-216 </td>
 <td></td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>VWR Scientific</td>
 <td>BDH4534-500GP</td>
 <td></td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>VWR Scientific</td>
 <td>BDH4532-500GP</td>
 <td></td>
 </tr>
 <tr>
 <td>MgSO4</td>
 <td>VWR Scientific</td>
 <td>BDH0246-500GP</td>
 <td></td>
 </tr>
 <tr>
 <td>Ca(NO3)2</td>
 <td>VWR Scientific</td>
 <td>BDH0226-500GP</td>
 <td></td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>VWR Scientific</td>
 <td>BDH4520-500GP</td>
 <td></td>
 </tr>
 <tr>
 <td>Morpholinos</td>
 <td>GeneTools, LLC</td>
 <td></td>
 <td></td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae at JoVE.com

Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

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 ...

non-invasive-imaging-of-disseminated-candidiasis-in-zebrafish-larvae

Research

JoVE Journal

Immunology and Infection

14.6K Views.  University of Maine. 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.

Video: Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists. The value of 2P microscopy is that it provides rich spatiotemporal information regarding cell behaviors within intact tissues and in live mice. Leukocyte recruitment plays a significant role in host defense against infection and when unchecked, can contribute to inflammatory and autoimmune diseases. Studying leukocyte recruitment in vivo is technically challenging since cells are moving rapidly within vessels located deep within light scattering tissues. To date, most intravital imaging studies require surgical preparation to expose the blood vessels and tissues. To avoid the tissue damage and inflammation induced by surgery itself, here, we describe a non-invasive single-cell imaging approach that can be used to study leukocyte trafficking in the mouse footpad and phalanges. We discuss the technical aspects of our 2P imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Here, we describe a non-invasive two-photon (2P) microscopy approach to study leukocyte homing in the mouse footpad. We discuss the technical aspects of our tissue imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists.  The value of 2P microscopy is that it ...

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal features of the natural infection. This protocol describes procedures for the establishment and evaluation of systemic infection due to neuropathogenic Escherichia coli K1 in the neonatal rat. Colonization of the gastrointestinal tract leads to dissemination of the pathogen along the gut-lymph-blood-brain course of infection and the model displays strong age dependency. A strain of E. coli O18:K1 with enhanced virulence for the neonatal rat produces exceptionally high rates of colonization, translocation to the blood compartment and invasion of the meninges following transit through the choroid plexus. As in the human host, penetration of the central nervous system is accompanied by local inflammation and an invariably lethal outcome. The model is of proven utility for studies of the mechanism of pathogenesis, for evaluation of therapeutic interventions and for assessment of bacterial virulence.

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Here, a procedure is described for the establishment of systemic infection in the neonatal rat with cultures of Escherichia coli K1. This non-invasive procedure permits colonization of the gastrointestinal tract, translocation of the pathogen to the systemic circulation, and invasion of the central nervous system at the choroid plexus.

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal ...

Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and pneumothorax from pleural disruption can result in a significant mortality rate. This protocol is a simple method of rat thymectomy that utilizes a mini-sternotomy and endotracheal intubation. Intubation is accomplished with a non-invasive and easily reproducible method and allows for positive pressure ventilation to prevent pneumothorax and a controlled airway that allows sufficient time for careful thymus dissection to minimize pleural disruption. A 1.5 cm sternal incision decreases contact with mediastinal vessels and pleura, while still providing full visualization of the thymus. Following exposure of the mediastinum, the thymus is removed by blunt dissection under magnification. The pleural space is then sealed by suture closure of the pre-tracheal muscles followed by the application of surgical glue. The thorax is then closed by suture closure of the sternum, followed by suture closure of the skin. All thymectomies were complete as evidenced by immunohistochemical (IHC) staining of mediastinal tissue, and absence of na&#239;ve T-cells by flow cytometry, and the procedure had a 96% survival rate. This method is suitable when complete thymectomy with minimal complications is desired for further immunological studies in athymic adult rats.

Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described.

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and ...

Modeling Mucosal Candidiasis in Larval Zebrafish by Swimbladder Injection

Early defense against mucosal pathogens consists of both an epithelial barrier and innate immune cells. The immunocompetency of both, and their intercommunication, are paramount for the protection against infections. The interactions of epithelial and innate immune cells with a pathogen are best investigated in vivo, where complex behavior unfolds over time and space. However, existing models do not allow for easy spatio-temporal imaging of the battle with pathogens at the mucosal level.
The model developed here creates a mucosal infection by direct injection of the fungal pathogen, Candida albicans, into the swimbladder of juvenile zebrafish. The resulting infection enables high-resolution imaging of epithelial and innate immune cell behavior throughout the development of mucosal disease. The versatility of this method allows for interrogation of the host to probe the detailed sequence of immune events leading to phagocyte recruitment and to examine the roles of particular cell types and molecular pathways in protection. In addition, the behavior of the pathogen as a function of immune attack can be imaged simultaneously by using fluorescent protein-expressing C. albicans. Increased spatial resolution of the host-pathogen interaction is also possible using the described rapid swimbladder dissection technique.
The mucosal infection model described here is straightforward and highly reproducible, making it a valuable tool for the study of mucosal candidiasis. This system may also be broadly translatable to other mucosal pathogens such as mycobacterial, bacterial or viral microbes that normally infect through epithelial surfaces.

In vivo spatio-temporal interactions of pathogen and immune defenses at the mucosal level are not easily imaged in existing vertebrate hosts. The method presented here describes a versatile platform to study mucosal candidiasis in live vertebrates using the swimbladder of the juvenile zebrafish as an infection site.

Early defense against mucosal pathogens consists of both an epithelial barrier and innate immune cells. The immunocompetency of both, and their ...

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of causing systemic disease in both fish and humans. A better understanding of S. iniae disease pathogenesis requires an appropriate model system. The genetic tractability and the optical transparency of the early developmental stages of zebrafish allow for the generation and non-invasive imaging of transgenic lines with fluorescently tagged immune cells. The adaptive immune system is not fully functional until several weeks post fertilization, but zebrafish larvae have a conserved vertebrate innate immune system with both neutrophils and macrophages. Thus, the generation of a larval infection model allows the study of the specific contribution of innate immunity in controlling S. iniae infection.
The site of microinjection will determine whether an infection is systemic or initially localized. Here, we present our protocols for otic vesicle injection of zebrafish aged 2-3 days post fertilization as well as our techniques for fluorescent confocal imaging of infection. A localized infection site allows observation of initial microbe invasion, recruitment of host cells and dissemination of infection. Our findings using the zebrafish larval model of S. iniae infection indicate that zebrafish can be used to examine the differing contributions of host neutrophils and macrophages in localized bacterial infections. In addition, we describe how photolabeling of immune cells can be used to track individual host cell fate during the course of infection.

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

Here, we present a protocol for the generation and imaging of a localized bacterial infection in the zebrafish otic vesicle.

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of ...

Deciphering and Imaging Pathogenesis and Cording of Mycobacterium abscessus in Zebrafish Embryos

Zebrafish (Danio rerio) embryos are increasingly used as an infection model to study the function of the vertebrate innate immune system in host-pathogen interactions.&#160;The ease of obtaining large numbers of embryos, their accessibility due to external development, their optical transparency as well as the availability of a wide panoply of genetic/immunological tools and transgenic reporter line collections, contribute to the versatility of this model.&#160;In this respect, the present manuscript describes the use of zebrafish as an&#160;in vivo&#160;model system to investigate the chronology of Mycobacterium abscessus infection. This human pathogen can exist either as smooth (S) or rough (R) variants, depending on cell wall composition, and their respective virulence can be imaged and compared in zebrafish embryos and larvae. Micro-injection of either S or R fluorescent variants directly in the blood circulation via the caudal vein, leads to chronic or acute/lethal infections, respectively. This biological system allows high resolution visualization and analysis of the role of mycobacterial cording in promoting abscess formation. In addition, the use of fluorescent bacteria along with transgenic zebrafish lines harbouring fluorescent macrophages produces a unique opportunity for multi-color imaging of the host-pathogen interactions. This article describes detailed protocols for the preparation of homogenous M. abscessus inoculum and for intravenous injection of zebrafish embryos for subsequent fluorescence imaging of the interaction with macrophages. These techniques open the avenue to future investigations involving mutants defective in cord formation and are dedicated to understand how this impacts on M. abscessus pathogenicity in a whole vertebrate.

Deciphering and Imaging Pathogenesis and Cording of Mycobacterium abscessus in Zebrafish Embryos

Optically transparent zebrafish embryos are widely used to study and visualize in real time the interactions between pathogenic microorganisms and the innate immune cells. Micro-injection of Mycobacterium abscessus, combined with fluorescence imaging, is used to scrutinize essential pathogenic features such as cord formation in zebrafish embryos.

Zebrafish (Danio rerio) embryos are increasingly used as an infection model to study the function of the vertebrate innate immune system in host-pathogen ...

A Non-invasive and Technically Non-intensive Method for Induction and Phenotyping of Experimental Bacterial Pneumonia in Mice

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant preclinical advances in our understanding of the underlying cellular and molecular pathogenesis. In vivo mouse models capture the integrated physiology and resilience of the host defense response in a manner not revealed by alternative, simplified ex vivo approaches. Several methods have been described in the literature for intrapulmonary inoculation of bacteria in mice, including aerosolization, intranasal delivery, peroral endotracheal cannulation under &#39;blind&#39; and visualized conditions, and transcutaneous endotracheal cannulation. All methods have relative merits and limitations. Herein, we describe in detail a non-invasive, technically non-intensive, inexpensive, and rapid method for intratracheal delivery of bacteria that involves aspiration (i.e., inhalation) by the mouse of an infectious inoculum pipetted into the oropharynx while under general anesthesia. This method can be used for pulmonary delivery of a wide variety of non-caustic biological and chemical agents, and is relatively easy to learn, even for laboratories with minimal prior experience with pulmonary procedures. In addition to describing the aspiration pneumonia method, we also provide step-by-step procedures for assaying the subsequent in vivo pulmonary innate immune response of the mouse, in particular, methods for quantifying bacterial clearance and the cellular immune response of the infected airway. This integrated and simple approach to pneumonia assessment allows for rapid and robust evaluation of the effect of genetic and environmental manipulations upon pulmonary innate immunity.

Several methods have been described in the literature for modeling bacterial pneumonia in mice. Herein, we describe a non-invasive, inexpensive, rapid method for inducing pneumonia via aspiration (i.e., inhalation) of a bacterial inoculum pipetted into the oropharynx. Downstream methods for assessment of the pulmonary innate immune response are also detailed.

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant ...

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo fluorescence optical imaging (OI), in two different mouse models of inflammation: a rheumatoid arthritis (RA) and a contact hypersensitivity reaction (CHR) model. Light with a wavelength in the near infrared (NIR) window (650 - 950 nm) allows a deeper tissue penetration and minimal signal absorption compared to wavelengths below 650 nm. The major advantages using fluorescence OI is that it is cheap, fast and easy to implement in different animal models.
Activatable fluorescent probes are optically silent in their inactivated states, but become highly fluorescent when activated by a protease. Activated MMPs lead to tissue destruction and play an important role for disease progression in delayed-type hypersensitivity reactions (DTHRs) such as RA and CHR. Furthermore, MMPs are the key proteases for cartilage and bone degradation and are induced by macrophages, fibroblasts and chondrocytes in response to pro-inflammatory cytokines. Here we use a probe that is activated by the key MMPs like MMP-2, -3, -9 and -13 and describe an imaging protocol for near infrared fluorescence OI of MMP activity in RA and control mice 6 days after disease induction as well as in mice with acute (1x challenge) and chronic (5x challenge) CHR on the right ear compared to healthy ears.

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper explains the application of fluorescent imaging using an activatable optical imaging probe to visualize the in vivo activity of key matrix metalloproteinases in two different experimental models of inflammation.

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo ...

A Non-invasive Way to Isolate and Phenotype Cells from the Conjunctiva

Traditionally, ocular surface cytology is studied with techniques such as spatula technology and brush technology. The problem with these techniques is that they may induce traumatic lesions on the surface of the eye, which can progress to scarring, eyelid deformity, limbal stem cell deficiency and in some cases, cause great discomfort to the subject. To avoid these clinical problems, impression cytology (IC) was developed to diagnose dry eye disease and later neoplasia, atopic disease, vernal keratoconjunctivitis and keratoconjunctivitis sicca. Typically, clinicians manually cut filter papers into required shapes and apply these to the ocular surface. Here, we describe how to perform IC using a commercially available medical device. This technique is explained here followed by immunophenotyping by flow cytometry. This technique requires less manual handling and causes less injury to the ocular surface.

The exposed normal ocular surface consists of cornea and conjunctiva. Epithelial cells, goblet cells and immune cells are present in the conjunctiva. Here, a non-invasive, technique of impression cytology is described using an impression cytology device and flow cytometry to analyze immune cells in the conjunctiva.

Traditionally, ocular surface cytology is studied with techniques such as spatula technology and brush technology. The problem with these techniques is ...

Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

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Chapters in this video

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Modeling Mucosal Candidiasis in Larval Zebrafish by Swimbladder Injection

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

Deciphering and Imaging Pathogenesis and Cording of Mycobacterium abscessus in Zebrafish Embryos

A Non-invasive and Technically Non-intensive Method for Induction and Phenotyping of Experimental Bacterial Pneumonia in Mice

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

A Non-invasive Way to Isolate and Phenotype Cells from the Conjunctiva