Authors greatly acknowledge the help of Bogdan Osadchiy and Evgenii Protasov (Irkutsk State University, Russia) in preparation of the video protocol. This research was supported by the Russian Science Foundation (#15-14-10008) and the Russian Foundation for Basic Research (#15-29-01003).

All experimental procedures were conducted in accordance with the EU Directive 2010/63/EU for animal experiments and have been approved by the Animal Subjects Research Committee of Institute of Biology at Irkutsk State University.
1. Fabrication of Microcapsules
NOTE: Microcapsules carrying a fluorescent dye are prepared using a layer-by-layer assembly of oppositely charged polyelectrolytes7,8. All procedures were performed at room temperature.
<ol>
	<li>To synthesize porous CaCO3 microcores enclosing the fluorescent dye, mix 2 mL of the SNARF-1-dextran solution (most polymer-bound fluorescent dye such as FITC-BSA can be used) at a concentration of ~2 mg/mL with 0.6 mL each of 1 mol/L solutions of CaCl2 and Na2CO3 under fast stirring. 
	NOTE: Pay attention to the different sensitivities of fluorescent dyes to photobleaching; if a light-sensitive fluorescent probe (like SNARF-1) is used, the manipulation and storage of the microparticles must be performed with as little light as possible.</li>
	<li>After 5-10 s of agitation, transfer the suspension to 2 mL microcentrifuge tubes and centrifuge for 15 s at 10,000-12,000 g to pellet CaCO3 microcores.</li>
	<li>Discard the supernatant, wash the cores with ~2 mL of deionized water, and resuspend the pellet by shaking.</li>
	<li>Repeat the centrifugation-washing procedure three times in total. After the last centrifugation, discard the supernatant.</li>
	<li>Incubate the microcores for 1 min in an ultrasonic bath to reduce their aggregation. 
	CAUTION: Do not forget to protect ears with headphones.</li>
	<li>To deposit the first polymeric layer on the templates, resuspend the cores in ~2 mL of a 4 mg/mL solution of poly(allylamine hydrochloride) (PAH) in 1 mol/L NaCl.
	<ol>
		<li>Keep the microcores in the solution for ~5 min with constant shaking.</li>
		<li>After 15 s of centrifugation, discard the supernatant with the unbound PAH. Wash the covered microcores with deionized water at least 3 times through multiple centrifugation and washing steps. After the last centrifugation, discard the supernatant.</li>
		<li>Incubate the microcores for 1 min in an ultrasonic bath to reduce their aggregation. 
		NOTE: If the applied fluorescent dye is cationic, start from poly(sodium 4-styrenesulfonate) (PSS) in 1 mol/L NaCl (see step 1.7).</li>
	</ol>
	</li>
	<li>Repeat step 1.6 with ~2 mL of a 4 mg/mL solution of PSS (also containing 1 mol/L NaCl) to deposit the second polymeric layer on the templates.</li>
	<li>Repeat steps 1.6 and 1.7 six times to deposit 12 polymeric layers. 
	NOTE: It is not recommended to take a long break (~12 h or more) in the procedure until ~3-5 layers have been deposited because CaCO3 microcores without coverage may tend to recrystallize. Note that PSS coverage causes a higher aggregation of the microcores, and the long pause is advisable only when PAH or poly-L-lysine grafted with polyethylene glycol (PLL-g-PEG) is the outmost layer.</li>
	<li>Incubate the covered microcores in 2 mg/mL PLL-g-PEG (~1 mL per microtube) for at least 2 h.
	<ol>
		<li>Wash the microcores with water via sequential centrifugation and resuspension steps. After the last centrifugation, discard the supernatant.</li>
	</ol>
	</li>
	<li>To obtain hollow microcapsules, dissolve the CaCO3 templates by adding 2 mL of 0.1 mol/L ethylenediaminetetraacetic acid (EDTA) solution (adjusted to pH 7.1 with NaOH) to the covered microcores.
	<ol>
		<li>After ~5 min of incubation, centrifuge the microcapsules for 45 s and discard the supernatant with the EDTA.</li>
		<li>Repeat steps 1.10-1.10.1 twice.</li>
	</ol>
	</li>
	<li>Wash the microcapsules with 0.9% NaCl three times through multiple centrifugation steps within 45 s followed by washing steps. After the last centrifugation step, discard the supernatant. 
	NOTE: The final microcapsule solution for injection must be kept sterile (for example by adding ampicillin, 0.1 mg/mL), and the media should be biocompatible with the object of investigation (isotonic media with neutral pH).</li>
	<li>Estimate the concentration of the prepared microcapsules in a hemocytometer under a fluorescence microscope. Take a series of pictures of the microcapsules, measure the diameter of about a hundred microcapsules using ImageJ9 or equivalent software, and investigate the size distribution using a histogram.</li>
	<li>Store the obtained encapsulated probe in the dark. 
	NOTE: After several washings in sterile 0.9% NaCl, the microcapsules can be stored for months at 4 &#176;C. Complete drying of the microcapsules during storage is not recommended.</li>
</ol>
2. Preparation of Optical Setup and Calibration of Microencapsulated SNARF-1
Note: Rough pH measurements with microencapsulated SNARF-1 can be made using images in two channels of a fluorescent microscope7, but in this protocol a one-channel fluorescent microscope connected to a fiber spectrometer was applied.
<ol>
	<li>Place the required set of fluorescence filters to the fluorescent microscope according to the characteristics of the applied fluorescent dye and turn on the fluorescent lamp.
	<ol>
		<li>Pull out the lever to the eyepieces. 
		CAUTION: Excess light can damage the spectrometer matrix. Thus, make sure that the lever is in the &#34;eyepiece&#34; mode when the spectrometer is not used.</li>
		<li>Connect one end of the optical fiber to the spectrometer and the other end to a collimator. Using adapters, place the collimator in the focus of the camera tube or other available port of the fluorescent microscope.</li>
		<li>Turn on the spectrometer. Run the spectrometer control program and prepare the spectrometer for measurements.</li>
	</ol>
	</li>
	<li>For calibration of the microcapsule batch, place ~5 &#181;L of the microcapsule suspension (~10 000 microcapsules per &#181;L in deionized water) on a microscope slide, and dry the drop in a dark place (for example, in a thermostat at 35 &#176;C).
	<ol>
		<li>To calibrate the spectral characteristics of the microencapsulated SNARF-1, use a series of buffers with different pH values in the range ~6-9. Drop ~10 &#181;L of a buffer onto the dried microcapsules with SNARF-1-dextran and cover it with a coverslip.</li>
		<li>Place the glass slide on the microscope stage. Locate the microcapsules using a &#215;40 objective.</li>
		<li>Turn the microscope lever to the camera port. Register their fluorescence with the spectrometer.Turn the lever back to the eyepieces. 
		NOTE: Make sure the spectral signal is far beyond the background level, and ensure that the microcapsules in the field of view are not in a bubble (by switching to a lower magnification if necessary). Avoid prolonged illumination of the same microcapsules. SNARF-1 is sensitive to photobleaching.</li>
		<li>Repeat step 2.2.3 for different microcapsules 10-15 times.</li>
	</ol>
	</li>
	<li>Calculate the fluorescence peak ratios (for example, using R or Scilab) for all registered spectra and determine the regression line between the median ratio (for each buffer) and medium pH using the following formula: 
	<img alt="Equation 1" src="/files/ftp_upload/57491/57491eq1.jpg" /> 
	NOTE: SNARF-1 has a spectrum with two peaks corresponding to the emission of the protonated and deprotonated dye, and the ratio between the peaks is responsive to the pH of the medium. In this study, the ratio between the fluorescence intensity for 605 and 660 nm is used. These wavelengths are chosen depending on the filter set used. a and b are coefficients to be determined by non-linear regression (for example, using R). Values 0.15 and 1.1 are, respectively, the minimum and maximum values of I605/I660 observed during the calibration.</li>
	<li>Collect about 10 &#181;L of fish blood from approximately 5 adult animals. Place fish into a Petri dish with 1 &#181;L/mL water suspension of clove oil for anesthesia and wait until the animal turns on its side and stops responding to pinches of the fin (usually ~2-3 min). Transfer the fish on a glass slide. Cut off the fish tail with a lancet and collect approximately 2 &#181;L of fish blood from the tail vein. 
	NOTE: To prevent blood clotting, treat the incision with heparin (5000 U/mL) and use heparinized glass capillaries and microcentrifuge tubes to collect the blood.
	<ol>
		<li>Drip about 10 &#181;L of blood with a pipette onto the tip of the microelectrode and determine the pH using a pH-meter.</li>
		<li>Drop the blood onto a slide with dried microcapsules and register the ratio of the fluorescence intensity as described for the calibration buffers (steps 2.2 - 2.3).</li>
	</ol>
	</li>
	<li>Adjust the linear coefficient of the calibration curve to make the curve match the measurements in the fish blood (for more details see Borvinskaya et al. 20175).</li>
</ol>
3. Preparation for Injection
<ol>
	<li>Release the steel needle from the tip of the insulin pen (or syringe) by removing the plastic with a sharp lancet. 
	NOTE: Any thin needle (&#216;0.33 mm or less) or glass capillary (usually &#216;1 mm) can be prepared for microinjection10,11.</li>
	<li>Insert the needle halfway into the glass microcapillary; quickly and gently solder it using a gas torch.</li>
	<li>Connect the glass microcapillary to the microinjector and flush it with sterile water three times. Ensure that the liquid flows through the needle.</li>
	<li>Fill the system with distilled water. 
	NOTE: Make sure there are no bubbles in the system.</li>
</ol>
4. Injection
<ol>
	<li>Resuspend the prepared suspension of microcapsules (in sterile 0.9% NaCl, or any other media used for the injections, with a concentration of 0.5 to 6 million microcapsules per microliter) using the ultrasonic bath for 1 min. 
	NOTE: Since the microcapsules tend to precipitate, during the following injection, shake the vial with the microcapsules mechanically (using a rotor) or manually every few minutes to resuspend them and prevent their aggregation.</li>
	<li>Place the fish into a Petri dish with anesthetic (0.1 mL/L of clove oil suspended in water) for ~2-3 min. Wait until the fish turns on its side and stops responding to a light pinch of the fin.</li>
	<li>Using a spoon, transfer the fish out of the anesthetic and gently place it on a damp sponge in a lateral position with the head towards the left (for right-handed person) or towards the right (for left-handed person).</li>
	<li>Just before the injection, suck 1-2 mm of air into the glass capillary connected with microinjector. Then, load it with approximately 2 &#181;L of the dispersed microcapsules. 
	NOTE: Before the injection, the microcapsule solution must be adjusted to the temperature at which the fish are kept.</li>
	<li>Gently stabilize the body of the fish on the sponge with non-dominant hand.
	<ol>
		<li>Find the lateral line of the fish. Mentally select a segment that extends from the operculum to the end of the abdominal cavity. Find the middle of this segment. Put the needle 1 mm lower in the ventral direction.</li>
		<li>With a scraping movement, gently move the fish scales aside, and make a puncture. Insert the needle into the body at an angle of 45&#176; to the table surface.</li>
		<li>Push the needle toward the spine until it carefully rests against it.</li>
		<li>Release approximately 1 &#181;L of the microcapsules&#39; suspension into the kidney, and slowly withdraw the needle. 
		NOTE: To find the proper puncture site more easily, it is useful to practice finding the trunk kidney by transilluminating the fishes using a bottom light, as shown in Figure&#160;2A and 2B.</li>
	</ol>
	</li>
	<li>Rinse the fish from head to tail with a stream of water to remove any spilled microcapsules at the injection site.</li>
</ol>
5. In Vivo Visualization
<ol>
	<li>Use the dissection scissors to remove the gill cover from the fish head and denude the fish gills. Rinse the gills with water.</li>
	<li>Using a spoon, transfer the fish to a microscope slide, and place it on the stage of the fluorescent microscope. 
	NOTE: Make sure that the gills of the fish do not dry out during successive procedures. To avoid this, periodically moisten them with water using a Pasteur pipette (approximately every 1-2 min).</li>
	<li>Darken the room and using low magnification (x10&#160;objective) inspect the gills to find the fluorescent microcapsules. 
	NOTE: When the procedure is used for introduction of some fluorescent particles into fish circulatory system, it is recommended to inspect gills of several individuals for unexpected fluorescent particles before injections. Gills of wild-type zebrafish do not have autofluorescence, but in some cases sporadic fluorescent particles (like food pieces or unicellular symbionts) may be present on gills. If necessary, such particles can be recognized based on their specific shape (for example, food pieces have irregular shape, unlike spherical microcapsules) or fluorescence spectrum (i.e., color).
	<ol>
		<li>Switch the lens to a higher magnitude (&#215;40 objective), and position a microcapsule or a group of microcapsules in the center of the field of view.</li>
		<li>Turn the lever to the port with a connected spectrometer. Record the spectral signal.</li>
		<li>Turn the lever back to the eyepiece.</li>
		<li>Repeat the measurements for different microcapsules several times.</li>
	</ol>
	</li>
	<li>Transfer the fish to the aquarium with proper aeration for recovery. 
	NOTE: With minimal practice, it is possible to perform the injection and signal recording at an approximate rate of 2-3 min per fish. The measurement can be repeated for one individual several times with the use of repeated low, harmless doses of anesthesia or another method of fixation. For long-term observation, use a system with continuous anesthesia12.</li>
</ol>

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The authors have nothing to disclose.

To demonstrate the injection of microparticles into the zebrafish kidney, we used semi-permeable microcapsules loaded with an indicator dye. Thus, the protocol contains instructions for the fabrication of the microcapsules using the layer-by-layer assembly of oppositely charged polyelectrolytes7,8,15,16,17,18 (Figure 1A). An advantage of this technology is that it is easy to perform with available laboratory equipment. Depending on the conditions and compounds used, the fabricated microcapsules can range from 0.5 to 100 &#181;m with nanometers-thick polyelectrolyte shell15. The synthesis parameters described in this manuscript result in elastic microcapsules consisting of 12 layers (in addition to the final biocompatible layer) of polymers approximately 2-6 &#181;m in size (Figure&#160;1B and 1C). The most important step determining the size of the microcapsules is the formation of the template microcores. This process involves the spontaneous formation of calcium carbonate crystals, and therefore, the obtained particles are non-uniform. Hence, a characterization of the microcapsule size distribution should be performed for every batch.
The important stage of this protocol is the optimization of the delivery of microparticles into the zebrafish circulatory system without the use of micromanipulation techniques. The administration of whole cells by retro-orbital injection has been previously described in detail3. However, in our experience, it is not easy for novice users to gain the injection efficiency described by the authors because the retro-orbital sinus is very small and located close to the pharynx and gill arches, which are easy to accidentally injure with a needle. Since less than a millimeter accuracy is required for the injections, the injecting properly is a rather difficult task. An equally quick and more efficient alternative (Table 1) is to inject directly into the fish kidney parenchyma. During the injection, the needle mechanically damages renal capillaries and blood vessels (e.g., the largest posterior cardinal vein), which allows entrance of microparticles into the circulatory system (Figure 2E). Finally, the central bulge of the trunk kidney in adult zebrafish is large enough (up to 2 mm) for a manual injection without microsurgical equipment.
The proper positioning of the injection needle is critical for a successful manual administration. You can practice finding the trunk kidney in intact animals by transilluminating fishes using a bottom light, as shown in Figure&#160;2A and 2B. The scheme in Figure 2B demonstrates how to properly perform the injection. If the procedure fails to administer microparticles into the blood stream, re-injection can be made at the same puncture within a short time frame with virtually no effect on the survival rate of the individuals.
Injection into the trunk kidney of adult zebrafish (also successfully tested on adult guppy Poecilia reticulata) is an effective method of microparticle delivery into the fish bloodstream with an allowable animal mortality; however, it is not perfectly suitable for drug administration due to variations in the injected volume. A weak point of this technique is a significant leakage of the solution out of the kidney into the abdomen because of the rapid administration. Nevertheless, a relative amount of the substance injected into bloodstream can be roughly estimated using visualization in the gill capillaries (Table 2). There is no strict limitation of the injection volume, but the administration of more than 1 &#181;L of the suspension appears to be ineffective. The finest glass capillaries can be applied for the microinjection; however, because a large number of microcapsules tends to incite aggregation, we recommend the use of 31-29G (&#216; 0.33-0.25 mm) steel needles to avoid clogs in the needle lumen.
The success of microcapsule delivery through kidney injection into the bloodstream should be monitored using the rapid inspection for the presence of the fluorescent particles in the gills. Gills are easily accessible organs for in vivo observations. The gill filaments are a network of blood capillaries covered by a thin layer of respiratory epithelium, which makes intravital imaging of the blood flow in the gills particularly convenient (a kind of natural optical window). To perform the observation, the cartilage gill cover can be cut to expose the filaments. This procedure is safe for fish, which can live with denuded gills in well aerated water without a decrease in life expectancy. Moreover, the gills of large fish can be examined under a microscope simply by pushing the operculum out of the way19. In case of a successful injection, fluorescent microparticles immediately appear in the gills (Figure 3). Note that implanted microparticles are considerably dissolved in the circulating blood volume, and a reduced number of particles reach the gill capillaries, thus sufficient concentration of microparticles should be selected for detection in fish gills after injection into fish kidney (Table 2).
The proposed procedure for implantation can be applied in a wide range of studies involving different species of small fishes. Despite the technique having been developed and optimized for injection of fluorescent microparticles into circulatory system, it can also be applied for implantation of non-colored micro/nanoparticles or dissolved substances; however, in this case the effectiveness of injection should be verified in some other way. The currently described steps are optimal for such purposes as physiological monitoring using different optical micro- or nanosensors, visualization of vasculature, delivery of vaccines or drugs on some optically visible carriers, and implantation of genetically modified cells.

The administration of micro-size particles into an animal organism is an important task in such areas as drug and vaccine delivery1, vasculature visualization2, transgenic cell implantation3, and tiny optical sensor implantation4,5. However, the implantation procedure for microscale particles into the vascular system of small laboratory animals is difficult, especially for delicate aquatic organisms. For popular research specimens like zebrafish, it is advised that these procedures be clarified using video protocols.
Intracardiac and capillary microinjections require trained personnel and unique microsurgery facilities for the delivery of microobjects into zebrafish blood. Previously, a retro-orbital manual injection3&#160;was suggested as an easy and effective method for the administration of whole cells. However, in our experience, because of the small area of the eye capillary network, it takes much practice to achieve the desired outcome from this technique.
Herein, we describe a method for robust and efficient microparticle implantation into the circulatory system by manual injection directly into the kidney tissue of adult zebrafish, which is rich in capillaries and renal vessels. This technique is based on the video protocol for cell transplantation into the zebrafish kidney6, but the traumatic and time-consuming microsurgical steps were eliminated. The proposed method is characterized by low mortality (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment.
An important part of the proposed protocol is the visualization of the implanted microparticles (if they are fluorescent or colorized) in the gill capillaries, which allows for the verification of the injection quality, a rough relative assessment of the number of injected particles, and the detection of the spectral signal for physiological measurements directly from the circulating blood. As an example of the possible applications of the described technique, we demonstrate the protocol for in vivo measurements of zebrafish blood pH using a microencapsulated fluorescent probe, SNARF-1, originally suggested in Borvinskaya et al. 20175.

<table><tbody><tr><td>SNARF-1-dextran, 70000 MW</td><td>Thermo Fisher Scientific</td><td>D3304</td><td>Fluorescent probe. Any other appropriate polymer-bound fluorescent dye can be used as a microcapsule filler</td></tr><tr><td>Albumin-fluorescein isothiocyanate conjugate (FITC-BSA)</td><td>SIGMA</td><td>A9771</td><td>Fluorescent probe</td></tr><tr><td>Rhodamine B isothiocyanate-Dextran (RITC-dextran)</td><td>SIGMA</td><td>R9379</td><td>Fluorescent probe</td></tr><tr><td>Calcium chloride</td><td>SIGMA</td><td>C1016</td><td>CaCO&lt;sub&gt;3&lt;/sub&gt; templates formation</td></tr><tr><td>Sodium carbonate</td><td>SIGMA</td><td>S7795</td><td>CaCO&lt;sub&gt;3&lt;/sub&gt; templates formation</td></tr><tr><td>Poly(allylamine hydrochloride), MW 50000 (PAH)</td><td>SIGMA</td><td>283215</td><td>Cationic polymer</td></tr><tr><td>Poly(sodium 4-styrenesulfonate), MW 70000 (PSS)</td><td>SIGMA</td><td>243051</td><td>Anionic polymer</td></tr><tr><td>Poly-L-lysine [20 kDa] grafted with polyethylene glycol [5 kDa], g = 3.0 to 4.5 (PLL-g-PEG)</td><td>SuSoS</td><td>PLL(20)-g[3.5]-PEG(5)</td><td>Final polymer to increase the biocompatibility of microcapsules</td></tr><tr><td>Sodium chloride</td><td>SIGMA</td><td>S8776</td><td>To dissolve applied polymers</td></tr><tr><td>Water Purification System</td><td>Millipore</td><td>SIMSV0000</td><td>To prepare deionized water</td></tr><tr><td>Magnetic stirrer</td><td>Stegler</td><td></td><td>For CaCO&lt;sub&gt;3&lt;/sub&gt; templates formation</td></tr><tr><td>Eppendorf&amp;nbsp;Research&amp;nbsp;plus pipette, 1000 &amp;micro;L</td><td>Eppendorf</td><td></td><td>Dosing solutions</td></tr><tr><td>Eppendorf&amp;nbsp;Research&amp;nbsp;plus pipette, 10 &amp;micro;L</td><td>Eppendorf</td><td></td><td>Dosing solutions</td></tr><tr><td>Pipette tips, volume range 200 to 1000 &amp;micro;L</td><td>F.L. Medical</td><td>28093</td><td>Dosing solutions</td></tr><tr><td>Pipette tips, volume range 0.1-10&amp;nbsp;&amp;mu;L</td><td>Eppendorf</td><td>Z640069</td><td>Dosing solutions</td></tr><tr><td>Mini-centrifuge Microspin 12, High-speed</td><td>BioSan</td><td></td><td>For microcapsule centrifugation-washing procedure</td></tr><tr><td>Microcentrifuge tubes, 2 mL</td><td>Eppendorf</td><td>Z666513</td><td>Microcapsule synthesis and storage</td></tr><tr><td>Shaker Intelli-mixer RM-1L</td><td>ELMY Ltd.</td><td></td><td>To reduce microcapsule aggregation</td></tr><tr><td>Ultrasonic cleaner</td><td></td><td></td><td>To reduce microcapsule aggregation</td></tr><tr><td>Head phones&amp;nbsp;</td><td></td><td></td><td>To protect ears from ultrasound</td></tr><tr><td>Ethylenediaminetetraacetic acid</td><td>SIGMA</td><td>EDS</td><td>To dissolve the CaCO&lt;sub&gt;3&lt;/sub&gt; templates</td></tr><tr><td>Monosodium phosphate</td><td>SIGMA</td><td>S9638</td><td>Preparation of pH buffers</td></tr><tr><td>Disodium phosphate</td><td>SIGMA</td><td>S9390</td><td>Preparation of pH buffers</td></tr><tr><td>Sodium hydroxide</td><td>SIGMA</td><td>S8045</td><td>To adjust the pH of the EDTA solution and buffers</td></tr><tr><td>Thermostat chamber</td><td></td><td></td><td>To dry microcapsules on glass slide</td></tr><tr><td>Hemocytometer blood cell count chamber</td><td></td><td></td><td>To investigate the size distribution and concentration of the prepared microcapsules</td></tr><tr><td>Fluorescent microscope Mikmed 2</td><td>LOMO</td><td></td><td>In vivo visualization of microcapsules in fish blood</td></tr><tr><td>Set of fluorescent filters for SNARF-1 (should be chosen depending on the microscope model; example is provided)</td><td>Chroma</td><td>79010</td><td>Visualization of microcapsules with fluorescent probes</td></tr><tr><td>Fiber spectrometer QE Pro</td><td>Ocean Optics</td><td></td><td>Calibration of microcapsules under microscope</td></tr><tr><td>Optical fiber QP400-2-VIS NIR, 400 &amp;mu;m, 2 m</td><td>Ocean Optics</td><td></td><td>To connect spectrometer with microscope port</td></tr><tr><td>Collimator F280SMA-A</td><td>Thorlabs</td><td></td><td>To connect spectrometer with microscope port</td></tr><tr><td>Glass microscope slide</td><td>Fisherbrand</td><td>12-550-A3</td><td>Calibration of microcapsules under microscope</td></tr><tr><td>Coverslips, 22 x 22 mm</td><td>Pearl</td><td>MS-SLIDCV</td><td>Calibration of microcapsules under microscope</td></tr><tr><td>Glass microcapillaries Intra MARK, 10 &amp;micro;L</td><td>Blaubrand</td><td>BR708709</td><td>To collect fish blood</td></tr><tr><td>Clove oil</td><td>SIGMA</td><td>C8392</td><td>Fish anesthesia</td></tr><tr><td>Lancet No 11</td><td>Apexmed international B.V.</td><td>P00588</td><td>To cut the fish tail and release the steel needle from the tip of insulin autoinjector</td></tr><tr><td>Heparin, 5000 U/mL</td><td>Calbiochem</td><td>L6510-BC</td><td>For treating all surfaces that come in contact with fish blood during fish blood collection</td></tr><tr><td>Seven 2 Go Pro pH-meter with a microelectrode</td><td>Mettler Toledo</td><td></td><td>To determine fish blood pH</td></tr><tr><td>Insulin pen needles Micro-Fine Plus, 0.25 x 5 mm</td><td>Becton, Dickinson and Company</td><td></td><td>For injection procedure. Any thin needle (&amp;Oslash; 0.33 mm or less) is appropriate</td></tr><tr><td>Glass capillaries, 1 x 75 mm</td><td>Hirschmann Laborger&amp;auml;te GmbH &amp;amp; Co</td><td>9201075</td><td>For injection procedure</td></tr><tr><td>Gas torch</td><td></td><td></td><td>To solder steel needle to glass capillary</td></tr><tr><td>Microinjector IM-9B</td><td>NARISHIGE</td><td></td><td>For precise dosing of microcapsules suspension</td></tr><tr><td>Petri dishes, 60 mm x 15 mm, polystyrene</td><td>SIGMA</td><td>P5481</td><td>For manipulations with fish under anesthesia</td></tr><tr><td>Plastic spoon</td><td></td><td></td><td>For manipulations with fish under anesthesia</td></tr><tr><td>Damp sponge</td><td></td><td></td><td>For manipulations with fish under anesthesia</td></tr><tr><td>Dissection scissors</td><td>Thermo Scientific</td><td>31212</td><td>To remove the gill cover from the fish head</td></tr><tr><td>Pasteur pipette, 3.5 mL</td><td>BRAND</td><td>Z331767</td><td>To moisten fish gills</td></tr></tbody></table>

simple and effective administration and visualization of microparticles in the circulatory system of small fishes using kidney injection

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, implantation of transgenic cells and tiny optical sensors. However, intravenous microinjections into small animals, which are mostly used in biological and veterinary laboratories, are very difficult and require trained personnel. Herein, we demonstrate a robust and efficient method for the introduction of microparticles into the circulatory system of adult zebrafish (Danio rerio) by injection into the fish kidney. To visualize the introduced microparticles in the vasculature, we propose a simple intravital imaging technique in fish gills. In vivo monitoring of the zebrafish blood pH was accomplished using an injected microencapsulated fluorescent probe, SNARF-1, to demonstrate one of the possible applications of the described technique. This article provides a detailed description of the encapsulation of pH-sensitive dye and demonstrates the principles of the quick injection and visualization of the obtained microcapsules for in vivo recording of the fluorescent signal. The proposed method of injection is characterized by a low mortality rate (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment. All described procedures can be performed on other small fish species, such as guppies and medaka.

The obtained results come from one of the three main categories of the presented protocol: the formation of fluorescent microparticles by encapsulation of a fluorescent dye (Figure 1), the kidney injection of microcapsules with further visualization in gill capillaries (Figure&#160;2&#160;and 3) and, finally, the in vivo spectral recording of SNARF-1 fluorescence to monitor blood pH levels (Figure 4).
The layer-by-layer approach using coating of the template CaCO3 cores enclosing the dye by multiple layers of oppositely charged polymers (PAH and PSS) and an outermost layer of a biocompatible polymer (PLL-g-PEG) is a simple and cost-effective method allowing to encapsulate different fluorescent probes such as SNARF-1-dextran, FITC-BSA or others (Figure 1A). As a result, hollow microcapsules of micrometric size with stable semi-permeable elastic shells and loaded with the fluorescent dye are obtained (Figure 1B). The microcapsules fabricated by this technique are typically non-uniform, with a normal distribution of particle size and characteristic median size in each batch (Figure 1&#1057;).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57491/57491fig1.jpg" /> 
Figure 1: Layer-by-layer&#160;synthesis and characterization of hollow polyelectrolyte microcapsules loaded with a fluorescent dye. (A) Scheme of a microcapsules assembly (drawn from a similar scheme published in Gurkov et al. 20164). (B) Picture of&#160;hollow microcapsules loaded with fluorescent dye FITC-BSA. (C) Size characterization of the batch of microcapsules from Figure 1B. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57491/57491fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Robust and efficient microparticle implantation into the circulatory system of small fishes can be performed by manual injection directly into the fish kidney. The puncture in the proper site of the fish body (trunk kidney) is crucial for rapid delivery of microparticles by manual injection. The fish kidney is a hematopoietic organ and contains pigmented melanophores, and thus, it is well colored. Because it is nestled between transparent chambers of the swim bladder, it is easy to identify the organ in the intact animal with weak pigmentation or by transillumination of small fishes using a bottom light source, as shown in Figure&#160;2A and 2B.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57491/57491fig2.jpg" /> 
Figure 2: Localization of the proper site for kidney injection. (A) Trans-illumination of the zebrafish demonstrates the localization of the trunk kidney (arrow). (B) Scheme illustrates how to identify the proper puncture site. The white dotted line indicates the lateral line of the abdominal segment of the fish. The arrow indicates the site for puncture and the proper direction of injection toward the spine. The swim bladder is designated by the green line. (C) Visualization of zebrafish kidney following the removal of organs and the body wall. An arrow points to the central bulge of the trunk kidney. (D) A sagittal histological section of D. rerio illustrates the general anatomy of the internal organs of the adult fish&#160;(H&#38;E stain). (E) Micrograph of a fish kidney (dotted) scaled from (D) with an arrow pointing to the lumen of the posterior cardinal vein. Asterisks indicate the swim bladder. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57491/57491fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To check the success of the injection procedure, a rapid visual inspection of the gills at low magnification (x10-20&#160;objective) should be made after cutting the fish gill cover (Figure 3A). Despite most of the microcapsules remaining at the injection site or spilling into the body cavity (Figure 3B), if the injection is performed correctly, it is possible to observe the fluorescent microcapsules freely floating in the gill capillaries of the denuded fish gills immediately after the injection (Figure 3C). If no fluorescent particles are detected in the gills, it is possible to repeat the injection in the same puncture. Successful delivery to the bloodstream should be obtained in approximately 70-90% of the total injected fish.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57491/57491fig3.jpg" /> 
Figure 3: Kidney injection of microcapsules with further visualization in gill capillaries.&#160;(A) The overall scheme of the microcapsule delivery into the zebrafish bloodstream. (B) Fluorescent image of a zebrafish after the injection of the microcapsules with green FITC-BSA dye (the puncture site is indicated by an arrow). (C) This picture of the gills of fish, scaled from (B), demonstrates the successful delivery of microcapsules by kidney injection (gills of wild-type zebrafish have no autofluorescence). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57491/57491fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
During the injection&#160;directly into the fish trunk kidney, extensive bruising normally forms beneath the puncture site, indicating damage to the parenchyma capillaries or the renal vessels. There is no blood leakage out of the body because the puncture appears to quickly contract. Despite internal bleeding, individuals can still recover with approximately 80-90% surviving through the procedure (Table 1). It is also known that fish species can regenerate nephrons de novo after injury13,14. If more than 20% of fish are dying, one must ensure that the anesthetization is performed properly.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="2">Encapsulated fluorescent dye</td>
			<td rowspan="2">n</td>
			<td rowspan="2">Concentration, microcapsules per &#181;L</td>
			<td rowspan="2">Injection volume, &#181;L</td>
			<td rowspan="2">Average diameter of microcapsules, &#181;m</td>
			<td rowspan="2">Successful delivery into the fish bloodstream, %</td>
			<td colspan="2">Mortality after injection, %</td>
		</tr>
		<tr>
			<td>1 h</td>
			<td>1 day</td>
		</tr>
		<tr>
			<td colspan="8">Injection into the trunk kidney</td>
		</tr>
		<tr>
			<td>FITC-BSA</td>
			<td>36</td>
			<td>4 x 106</td>
			<td>1.6</td>
			<td>2.7</td>
			<td>94</td>
			<td>8</td>
			<td>3</td>
		</tr>
		<tr>
			<td>SNARF-1-dextran</td>
			<td>29</td>
			<td>3-6 x 105</td>
			<td>1-2</td>
			<td>5.1</td>
			<td>72</td>
			<td>7</td>
			<td>12</td>
		</tr>
		<tr>
			<td>RITC-dextran</td>
			<td>20</td>
			<td>6 x 106</td>
			<td>2</td>
			<td>2.0</td>
			<td>70</td>
			<td>0</td>
			<td>0</td>
		</tr>
		<tr>
			<td>0.9% NaCl</td>
			<td>41</td>
			<td>-</td>
			<td>1-2</td>
			<td>-</td>
			<td>-</td>
			<td>5</td>
			<td>0</td>
		</tr>
		<tr>
			<td colspan="8">Retro-orbital injection</td>
		</tr>
		<tr>
			<td>FITC-BSA</td>
			<td>9</td>
			<td>1 x 105</td>
			<td>2</td>
			<td>4.2</td>
			<td>11</td>
			<td>22</td>
			<td>0</td>
		</tr>
		<tr>
			<td>RITC-dextran</td>
			<td>11</td>
			<td>6 x 106</td>
			<td>1</td>
			<td>2.0</td>
			<td>9</td>
			<td>18</td>
			<td>0</td>
		</tr>
	</tbody>
</table>
Table 1. Safety and efficiency of the delivery of microcapsules into the zebrafish bloodstream. The success of the procedure is determined by the presence of the fluorescent materials in the fish gill capillaries.
If the concentration of the injected microparticles is insufficient, too few particles may be available to be rapidly visualized in fish gills. At the same time, a suspension with a too high concentration can clog the needle. In case of layer-by-layer assembled microcapsules with median diameter ~2-5 &#181;m, a concentration of approximately 4*105-6*106 microcapsules per &#181;L is optimal for effortless detection in fish gills after injection into fish kidney (Table 2).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Concentration, microcapsules per &#181;L</td>
			<td colspan="7">Number of microcapsules in a microscope field of view (&#215;20 objective) in the gills of different individuals (n = 7 per concentration)</td>
		</tr>
		<tr>
			<td>4 x 106</td>
			<td>&#62; 100</td>
			<td>&#62; 100</td>
			<td>0</td>
			<td>&#62; 100</td>
			<td>&#8776; 70</td>
			<td>&#8776; 50</td>
			<td>&#62; 100</td>
		</tr>
		<tr>
			<td>4 x 105</td>
			<td>&#8776; 10</td>
			<td>&#8776; 20</td>
			<td>0</td>
			<td>0</td>
			<td>&#8776; 20</td>
			<td>&#8776; 40</td>
			<td>&#8776; 15</td>
		</tr>
		<tr>
			<td>4 x 104</td>
			<td>0</td>
			<td>3</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>4</td>
		</tr>
		<tr>
			<td>4 x 103</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
			<td>0</td>
		</tr>
	</tbody>
</table>
Table 2. Record of the visual counting of FITC-BSA-containing microcapsules in D. rerio gills after injection (1.6 &#181;L) into the trunk kidney.
The proposed method of microparticles&#39; implantation into fish circulatory system can be applied for in vivo monitoring of zebrafish blood pH by microencapsulated fluorescent probe, SNARF-1 (Figure 4). SNARF-1 has a spectrum with two peaks corresponding to the emission of the protonated and deprotonated dye (Figure 4A), thus the calibration curves of the ratio between the peaks at different pH of the media can be plotted (Figure 4B). The components of the blood influence the readout of the encapsulated SNARF-1 (Figure 4B), which can be evaluated experimentally by simultaneous measurement of blood pH by microcapsules and a pH-meter with a glass microelectrode. A putative calibration curve for microcapsules in zebrafish blood should be plotted by shifting the buffers&#39; curve by the coefficient equal to the pH difference measured experimentally (see Borvinskaya et al. 20175 for details).
Blood pH in zebrafish gill capillaries remains stable during at least several hours after the injection of microcapsules (Figure 4C). At the same time, a short hypercapnic exposure leads to a statistically significant decrease of blood pH, which demonstrates applicability of the method for in vivo research on small fishes.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57491/57491fig4.jpg" /> 
Figure 4: Representative example of monitoring of zebrafish blood &#1088;&#1053; by registration of the fluorescence spectra of microencapsulated dye SNARF-1.&#160;(A) Spectra of the prepared microcapsules loaded with SNARF-1 in sodium phosphate buffers at different pH. (B) Calibration curves of the prepared pH-sensitive microcapsules in sodium phosphate buffers and in extracted zebrafish blood. For all measurements, the mean &#177; standard deviation is depicted. (C) A representative example of the in vivo monitoring of zebrafish blood &#1088;&#1053; by encapsulated fluorescent SNARF-1 dye in zebrafish gill capillaries. In control conditions, blood pH remains stable during 4 hours after injection of microcapsules, while 5 minutes exposure under severe hypercapnia (900-1000 mg/L of dissolved CO2) causes acidification of the fish blood. Asterisk indicates statistically significant difference from the parallel control group with p &#60; 0.01 (Mann-Whitney U test).&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57491/57491fig4large.jpg">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection at JoVE.com

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, ...

simple-effective-administration-visualization-microparticles

Nanyang Technological University

Research

JoVE Journal

Biology

8.3K Views.  Institute of Biology at Irkutsk State University. This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

Video: Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

Transplantation of Whole Kidney Marrow in Adult Zebrafish

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an organism. The functional definition of a HSC is a transplanted cell that has the ability to reconstitute all the blood lineages of an irradiated recipient long term. This designation was established by decades of seminal work in mammalian systems. Using hematopoietic cell transplantation (HCT) and reverse genetic manipulations in the mouse, the underlying regulatory factors of HSC biology are beginning to be unveiled, but are still largely under-explored. Recently, the zebrafish has emerged as a powerful genetic model to study vertebrate hematopoiesis. Establishing HCT in zebrafish will allow scientists to utilize the large-scale genetic and chemical screening methodologies available in zebrafish to reveal novel mechanisms underlying HSC regulation. In this article, we demonstrate a method to perform HCT in adult zebrafish. We show the dissection and preparation of zebrafish whole kidney marrow, the site of adult hematopoiesis in the zebrafish, and the introduction of these donor cells into the circulation of irradiated recipient fish via intracardiac injection. Additionally, we describe the post-transplant care of fish in an "ICU" to increase their long-term health. In general, gentle care of the fish before, during, and after the transplant is critical to increase the number of fish that will survive more than one month following the procedure, which is essential for assessment of long term (&lt;3 month) engraftment. The experimental data used to establish this protocol will be published elsewhere. The establishment of this protocol will allow for the merger of large-scale zebrafish genetics and transplant biology.

In this article, we demonstrate a method to perform HCT in adult zebrafish.

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

Developmental Biology

Retro-orbital Injection in Adult Zebrafish

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is often the most effective and noninvasive form of delivery of an agent of interest. In the zebrafish (Danio rerio), IV injection of drugs has long been a challenge because of the small vessel diameter. This has also proved a significant hurdle for the injection of cells during hematopoeitic stem cell transplantation. Historically, injections into the bloodstream were done directly through the heart. However, this intra-cardiac procedure has a very high mortality rate as the heart is often punctured during injection leaving the fish prone to infection, massive blood loss or fatal organ damage. Drawing on our experience with the mouse, we have developed a new injection procedure in the zebrafish in which the injection site is behind the eye and into the retro-orbital venous sinus. This retro-orbital (RO) injection technique has been successfully employed in both the injection of drugs in the adult fish as well as transplantation of whole kidney marrow cells. RO injection has a much lower mortality rate than traditional intra-cardiac injection. Fish that are injected retro-orbitally tend to bleed less following injection and are at a much lower risk of injury to a major organ like the heart. Further, when performed properly, injected cells and/or drugs quickly enter the bloodstream allowing compounds to exert their effect on the whole fish and kidney cells to easily home to their niche. Thus, this new injection technique minimizes mortality while allowing efficient delivery of material into the bloodstream of adult fish. Here we exemplify this technique by retro-orbital injection of Tg(globin:GFP) cells into adult casper fish as well as injection of a red fluorescent dye (dextran, Texas Red ) into adult casper fish. We then visualize successful injections by whole animal fluorescence microscopy.

Here we show how to do retro-orbital injection in adult zebrafish.

Drug treatment of whole animals is an essential tool in any model system for pharmacological and chemical genetic studies. Intravenous (IV) injection is ...

Intravenous Microinjections of Zebrafish Larvae to Study Acute Kidney Injury

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections on 2 dpf zebrafish. This technique represents an efficient and rapid method to deliver soluble substances into the bloodstream of zebrafish larvae, allowing for the injection of 15-20 fish per hour. In addition to AKI studies, this microinjection technique can also be used for other types of experimental studies such as angiography.  We provide a detailed protocol of the technique from equipment required to visual measures of decreased kidney function. In addition, we also demonstrate the process of fixation, whole mount immunohistochemistry with a kidney tubule marker, plastic embedding and sectioning of the larval zebrafish. We demonstrate that zebrafish larvae injected with gentamicin show morphological features consistent with AKI: edema, loss of cell polarity in proximal tubular epithelial cells, and morphological disruption of the tubule.

We describe a technique of microinjecting the aminoglycoside, gentamicin, into 2 days post-fetilization (dpf) zebrafish larvae to induce acute kidney injury (AKI). We also describe a method for whole mount immunohistochemistry, plastic embedding and sectioning of zebrafish larvae to visualize the AKI mediated damage.

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections ...

Intramyocardial Injection for the Study of Cardiac Lymphatic Function in Zebrafish

Zebrafish have proved to be an important model for studying cardiovascular formation and function during postembryonic development and regeneration. The present protocol describes a method for injecting fluorescent tracers into the zebrafish myocardium to study interstitial fluid and debris uptake into cardiac lymphatic vessels. To do so, microspheres (200 nm diameter) and quantum dots (&#60;10 nm diameter) are introduced into the myocardium of live zebrafish, which can be tracked using ex vivo confocal microscopy. These tracers are then tracked intermittently over several hours to follow clearance from the myocardium into cardiac lymphatic vessels. Quantum dots&#160;are transported through cardiac lymphatic vessels away from the heart, while larger microspheres&#160;remain at the injection site for over three weeks. This method of intramyocardial injection can be extended to other uses, including the injection of encapsulated MS or hydrogels to locally release cells, proteins, or compounds of interest to a targeted region of the heart.

The present protocol allows efficient and stable delivery of fluorescent microspheres (MS) and quantum dots (QD) into the myocardium of live fish that can be tracked (traced) over time.

Zebrafish have proved to be an important model for studying cardiovascular formation and function during postembryonic development and regeneration. The ...

Transplantation of Cells Directly into the Kidney of Adult Zebrafish

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic diseases1. However, most organs are not easily accessible, necessitating the need to develop surgical methods to gain access to these structures. In this video article, we describe a method for transplanting cells directly into the kidney of adult zebrafish, a popular model to study regeneration and disease2. Recipient fish are pre-conditioned by irradiation to suppress the immune rejection of the injected cells3. We demonstrate how the head kidney can be exposed by a lateral incision in the flank of the fish, followed by the injection of cells directly in to the organ. Using fluorescently labeled whole kidney marrow cells comprising a mixed population of renal and hematopoietic precursors, we show that nephron progenitors can engraft and differentiate into new renal tissue - the gold standard of any cell-based regenerative therapy. This technique can be adapted to deliver purified stem or progenitor cells and/or small molecules to the kidney as well as other internal organs and further enhances the zebrafish as a versatile model to study regenerative medicine.

Cell transplantation is an essential technique for studying tissue regeneration and for developing cell-based therapies of disease. We demonstrate here a microsurgical technique that permits the transplantation of genetically labeled cells directly into the kidney of adult zebrafish fish.

Regenerative medicine based on the transplantation of stem or progenitor cells into damaged tissues has the potential to treat a wide range of chronic ...

Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish kidney develops and functions in almost the same way as humans. A major difference in the construction of the human kidney is the presence of millions of nephrons compared to the zebrafish that has only two. However, simplifying such a complex system into basic functional units has aided our understanding of how the kidney develops and operates. In zebrafish, the midline located glomerulus is responsible for the initial blood filtration into two pronephric tubules that diverge to run bilaterally down the embryonic axis before fusing to each other at the cloaca. The pronephric tubules are heavily populated by motile cilia that facilitate the movement of filtrate along the segmented tubule, allowing the exchange of various solutes before finally exiting via the cloaca2-4. Many genes responsible for CKD, including those related to ciliogenesis, have been studied in zebrafish5. However, a major draw back has been the difficulty in evaluating zebrafish kidney function after genetic manipulation. Traditional assays to measure kidney dysfunction in humans have proved non translational to zebrafish, mainly due to their aquatic environment and small size. For example, it is not physically possible to extract blood from embryonic staged fish for analysis of urea and creatinine content, as they are too small. In addition, zebrafish do not produce enough urine for testing on a simple proteinuria &#8216;dipstick&#8217;, which is often performed during initial patient examinations. We describe a fluorescent assay that utilizes the optical transparency of the zebrafish to quantitatively monitor the clearance of a fluorescent dye, over time, from the vasculature and out through the kidney, to give a read out of renal function1,6-9.

The zebrafish is a popular tool to model chronic kidney disease (CKD). However, their small size makes it impossible to evaluate renal function using traditional methods. We describe a fluorescent dye kidney clearance assay1 that allows quantitative analysis of zebrafish kidney function in CKD.

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...

Micropuncture of Bowman's Space in Mice Facilitated by 2 Photon Microscopy

Renal micropuncture and renal 2-photon imaging are seminal techniques in renal physiology. However, micropuncture is limited by dependence on conventional microscopy to surface nephron features, and 2-photon studies are limited in that interventions can only be assessed at the organ, rather than the nephron level. In particular, micropuncture studies of the glomeruli of mice have been challenged by the paucity of surface glomeruli in mice. To address this limitation in order to pursue studies of aspirate from Bowman&#39;s space in mouse physiologic models, we developed 2-photon glomerular micropuncture. We present a novel surgical preparation that allows lateral access to the kidney while preserving the required vertical imaging column for 2-photon microscopy. Administration of high molecular weight fluorescein isothiocyanate (FITC)-dextran is used to render the renal vasculature and therefore glomeruli visible for 2-photon imaging. A quantum dot-coated pipette is then introduced under stereotactic guidance to a glomerulus selected from the several to many which may be visualized within the imaging window. In this protocol, we provide details of the preparation, materials, and methods necessary to carry out the procedure. This technique facilitates previously-impossible physiologic study of the kidney, including recovery of filtrate from Bowman&#39;s space and all segments of the nephron within the imaging depth limit, about 100 &#181;m below the renal capsule. Pressure, charge and flow may all be measured using the introduced pipette. Here, we provide representative data from liquid chromatography/mass spectrometry performed on aspirate from Bowman&#39;s space. We expect this technique to have wide applicability in renal physiologic investigation.

We present use of 2-photon microscopy to place a micropipette within Bowman&#39;s urinary space in mice, combining 2 foundational techniques of renal physiology. Use of 2-photon microscopy overcomes critical limitations of conventional microscopy for micropuncture renal physiology studies.

Renal micropuncture and renal 2-photon imaging are seminal techniques in renal physiology. However, micropuncture is limited by dependence on conventional ...

Intrathoracic Injection for the Study of Adult Zebrafish Heart

The adult zebrafish heart provides a powerful model in cardiac regeneration research. Although the strength of this system is based on transgenic approaches, a rapid delivery of exogenous factors provides a complementary technique in functional studies. Here, we present a method that relies on administration of a few microliters of solution into the pericardial cavity without causing myocardial damage. Intrathoracic (IT) injections can efficiently deliver proteins and chemical compounds directly onto the heart surface. The injected substances diffuse through the epicardium into the underlying cardiac tissues. Compared to intraperitoneal (IP) injections, the main advantage of intrathoracic injections is the focal administration of the tested factors on the target organ. The delivery of molecules directly into the pericardium is a suitable strategy for studies of cardiac preconditioning and regeneration in adult zebrafish.

This method relies on the injection of 0.5&#8722;3 &#956;L of solution into the thorax of adult zebrafish. The procedure efficiently delivers proteins and chemical compounds into the proximity of the zebrafish heart without damaging the organ. The approach is suitable for testing effects of exogenous factors on various tissues of the heart.