Technical assistance provided by Jaipreet Bharj. This work was supported by grants from EU-FP7 (SYSCILIA -241955) and The Dutch Kidney Foundation (CP11.18).

Ethics Statement: Animal maintenance, husbandry, and procedures are defined and controlled by the Animals (Scientific Procedures) Act 1986. All animal experimentation has been carried out under licenses granted by the Home Secretary (PIL No. 70/7892) in compliance with Biological Services Management Group and the Biological Services Ethical Committee, SGUL, London, UK. All efforts were made to reduce the number of animals used and to refine both procedures and husbandry in order to minimize suffering and enhance welfare.
1. Preparation of Instruments, Anesthetic, and Fluorescent Dye
<ol>
	<li>Using a micropipette puller and borosilicate standard wall capillaries (without filaments) pull needles of appropriate length for microinjection (Figure 1A).</li>
	<li>Make an agarose mould for orientation of embryos for easy injection into the pericardium (Figure 1B). Glue approximately ten glass microscope slides together to form a &#8216;staircase&#8217; of offset slides. For best results, use a drop of rapid set epoxy resin glue in the center of each slide.</li>
	<li>Cut out a section of approximately 78 mm in length halfway through two 10 ml plastic pipettes using a Bunsen burner heated scalpel, remove pipette ends as appropriate. Attach and glue (epoxy resin) pipette sections to the stacked glass slides to provide stability to allow the mould to dip into a 90 mm Petri dish. Make every effort to ensure the slide corners align perpendicular to the Petri dish floor. Allow glue time to set.</li>
	<li>Cast the mould in a 2% agarose solution made in fish water (obtained from the aquarium). Pour agarose into a 90 mm Petri dish and place the mould on top of the open Petri dish, permitting the stacked slide edges to submerge at an angle under the agarose surface. Leave to set on the lab bench for 30 min.</li>
	<li>Remove the slide cast and cover the slide-imprinted agarose with fresh fish water containing Tricaine anesthetic (see step 1.9) at a 1:25 ratio.</li>
	<li>Make solutions of rhodamine dextran (RD) dye. Resuspend rhodamine B 10,000 MW labeled dextran in autoclaved ultrapure water to make a stock concentration of 50 mg/ml. For microinjection, further dilute the stock to a final concentration of 5 mg/ml in autoclaved ultrapure water. 
	NOTE: Pigmented cells begin to differentiate in zebrafish from 24 hr post fertilization (hpf); these can obscure fluorescent markers during image acquisition.</li>
	<li>Inhibit melanocyte formation by using N-Phenylthiourea (PTU), which blocks melanogenesis through inhibition of tyrosinase. Make a 0.003 % stock solution of PTU by dissolving PTU powder in aquarium water, heat solution at 60 &#176;C to fully solubilize.</li>
	<li>Incubate zebrafish embryos in methylene blue solution, a mild fungicide, to prevent fungal blooms and increase survival. Add 2 ml of methylene blue stock solution, containing 0.1% Methylene blue in ultrapure water, to 1 L of aquarium water and use as a standard embryo medium.</li>
	<li>Make up a 15 mM stock concentration of Tricaine/ethyl 3-aminobenzoate methanesulfonate salt as instructed in &#8216;The Zebrafish Book&#8217; 23. Anesthetize the embryos and consequently immobilized them to perform the microinjection procedure.</li>
	<li>After anesthetizing the embryos, orient them, for imaging by using the non-toxic and viscous properties of methylcellulose. To make a 200 ml preparation of 3% methylcellulose, chill 130 ml of water at -20 &#176;C for 30 min and place on ice. Heat 70 ml of water to 80 &#176;C in a glass beaker, add 6 g of methylcellulose, and agitate using a glass rod until all particles are wetted and evenly dispersed. Add the ice-cold water, mix, and allow the preparation to cool at 4 &#176;C for 30 min before aliquoting into 50 ml tubes. 
	NOTE: Lowering the temperature allows the methylcellulose to become soluble. The solution will become thicker as the powder hydrates. 3% methylcellulose can be stored without bacterial growth for short periods of a week at 4 &#176;C or longer at -20 &#176;C. Make sure the methylcellulose has reached RT before use.</li>
	<li>To perform injections of fluorescent dye into the zebrafish use a standard microinjection set-up (Figure 1C). This consists of an air compressor connected to a pressure regulator system that feeds into a straight pipette holder for use with 1.0 outer diameter capillaries. The pipette holder should be housed within an MM33 compact 3-axis control micromanipulator secured to a steel base plate by a magnetic stand. Embryos can be visualized, manipulated and injected using a stereo dissecting microscope.</li>
</ol>
2. Zebrafish Husbandry and Pre-injection Treatment
<ol>
	<li>Maintain zebrafish as previously described23. Use an aquarium set-up that provides recirculating water supplied at a constant temperature of 28.5 &#176;C, conditioned to pH 6.8 - 7.2 and conductivity of 450-550 &#956;S with sodium bicarbonate and Instant Ocean Sea Salt, respectively. Use wildtype or transgenic zebrafish lines as appropriate to the experiment, maintaining a stocking density of 15 males to 15 females per 8 L tank. Set the fish facility photocycle to 14 hr of daylight between 9 am - 11 pm.</li>
	<li>Zebrafish spawning commences at the beginning of the photocycle, when the lights turn on in the morning. To ensure maximum eggs can be collected without disturbing the fish, submerge breeding tanks (containing mesh inserts, available from zebrafish specialists) into wildtype stock tanks the evening before collection.</li>
	<li>On the day of collection, allow 30 - 40 min for the fish to spawn, after which collect and rinse eggs using a tea strainer and fresh aquarium water. Deposit the cleaned eggs in a 90 mm Petri dish containing embryo medium and incubate at 28.5 &#176;C.</li>
	<li>Treat the embryos with PTU to inhibit melanogenesis. At 8 hpf, transfer viable embryos in minimal liquid to fresh petri dishes containing 1:100 PTU to embryo medium and further incubate at 28.5 &#176;C until 72 hpf. 
	NOTE: PTU can cause developmental defects if used at earlier stages so should be restricted to post 8 hpf stages, but can be treated as late as 24 hpf. Late addition of PTU does not fully block eye pigmentation but is generally successful and inhibiting trunk melanocyte formation.</li>
</ol>
3. Microinjection
NOTE: The pericardium encases the heart but is separated for protection and ease of cardiac movement by fluid within the pericardial cavity. The aim of this procedure is to inject RD into the pericardial cavity, this allows for rapid uptake of the dye to the vasculature system.
<ol>
	<li>Load needles with 6 &#956;l - diluted RD, using microloader tips and secure into the needle holder of the micromanipulator. Break the end of the needle using fine tipped forceps (Figure 1A). Move the RD solution to the tip of the needle by maximizing pulse duration to the minute setting, switch back to msec once complete.</li>
	<li>Use a stage micrometer, the pressure regulator, and refinements to needle tip length (using forceps) to adjust the size of the expelled droplet to 100 &#956;m in diameter, this equates to a 0.5 nl volume (Figure 2A). 
	NOTE: When breaking the needle, be careful not to break too much off otherwise it will make calibrating the injection volume difficult. If necessary, further break the needle tip to increase the drop size however, maintain a needle thickness that permits entry into the pericardium without excessive bending or damage to the tissue.</li>
	<li>Prior to injection, anesthetize the embryos in embryo medium containing Tricaine. Set up 2 x 35 mm Petri dishes containing 5 ml embryo medium, label one &#8216;Tricaine&#8217; and the other &#8216;Recovery&#8217;.</li>
	<li>Add 200 &#956;l of stock Tricaine to the appropriately labeled dish. Once ready to inject, select an individual embryo at 72 hpf and transfer in minimal liquid to the Tricaine dish. Monitor the activity of the embryo, test the embryo is anesthetized and immobilized by gentle agitation using a truncated microloader tip.</li>
	<li>Transfer the anesthetized embryo to the injection mould and orientate the embryo within an agarose trough so the left side is facing up, positioning the heart to the left of the field of view.</li>
	<li>To inject the RD into the heart, pierce the pericardium with the needle and inject 1 nl of RD into the pericardial cavity of the anaesthetized embryo (Figure 2B, right panel). Withdraw the needle after injection. Transfer the injected embryo in minimal liquid to the &#8216;Recovery dish&#8217; and monitor for 1 min. Transfer the individual embryo to a 24-well plate containing 1 ml fresh embryo medium (containing PTU) and label appropriately. 
	NOTE: The pericardium is tough, however there is a notable weak point at the intersection where the pericardium meets the ventro-caudal pharyngeal arches and dorso-rostral yolk sac (Figure 2B arrowhead). The needle should be directed to this groove. When the needle is in place, a firm tap of a finger on top of the micromanipulator will facilitate entry into the pericardial cavity.</li>
	<li>Repeat steps 3.2 - 3.6 for at least 10 embryos per experimental group, place each fish in a separate well and uniquely labelto permit individual experimental follow-ups. Incubate at 28.5 &#176;C for 3 hr.</li>
</ol>
4. Imaging Acquisition
<ol>
	<li>At 3 hr post injection (hpi) anesthetize embryos as previously described and transfer to a 35 mm Petri dish containing 3% methylcellulose. Gently push embryos into the methylcellulose and orientate so the lateral side can be imaged.</li>
	<li>Acquire an image of the embryo under UV, using a filter for visualization of emitted light at 570 nm (Figure 2C). After image acquisition, allow the embryo to recover before replacing in the designated well. Acquire images for all injected embryos and further incubate at 28.5 &#176;C. 
	NOTE: In this protocol we used a fluorescent stereomicroscope with a TXR filter set, a DFC300FX camera and respective application software. Make a note of the exact acquisition settings for subsequent image acquisition.</li>
	<li>At 24 hpi, acquire a second round of images as described above. Once all the images have been acquired, at both 3 hpi and 24 hpi, humanely dispose of the embryos or use for other experimental means e.g., immunohistochemistry.</li>
</ol>
5. Image Processing
<ol>
	<li>Quantify fluorescent intensity of each injected embryo, at 3 hpi and 24 hpi, by analyzing images using NIH&#8217;s ImageJ software. To measure fluorescent intensity, open an image in ImageJ and specify a region of interest (roi) at 100 px2 (Edit&#160;&#8594; selection &#8594; specify).</li>
	<li>Position the heart in the center of the roi (Figure 2C), set measurements to include mean gray scale and area (Analyze &#8594; set measurements), perform measurement (Analyze &#8594; measure). Complete for each set of images, at 3 hpi and 24 hpi, per embryo and transfer average gray scale values to a spreadsheet for further processing. 
	NOTE: We selected a fixed roi size of 100 px2 as this encompasses individual hearts between embryos. The heart was selected to perform measurements due to its large size that enables a convenient location to measure fluorescent content of the blood prior to entering the kidney.</li>
	<li>Perform statistical analysis using appropriate statistical software. Compare groups for statistical significance using a student&#8217;s t-test.</li>
</ol>

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The authors have nothing to disclose.

Zebrafish offer a valuable tool to model human genetic disease, their use as a scientific instrument for&#160;in vivo&#160;research have enabled detailed studies of the genetic breakdown of many biological systems, including the kidney. Much is now understood about how the zebrafish kidney develops and functions. The striking similarities to human nephrogenesis and homology with disease causing genes21 has illustrated how zebrafish have become fundamental in understanding how defects in gene function ...

The human kidney plays a crucial role in filtering metabolic waste from the blood and recovering required solutes to sustain cellular homeostasis. There are a number of human genetic diseases that cause kidney dysfunction. The most common inherited renal disease is autosomal dominant polycystic kidney disease (ADPKD) characterized by the development of fluid filled sacs within nephritic tubules; the damage caused by cystogenesis is detrimental to kidney function10. ADPKD has an occurrence of 1:800 - 1:1,000 and accounts for 8 - 10% of patients in end stage renal failure (ESRF)11. Several genes have been implicated to cause ADPKD including polycystin-1 (PKD1) and -2 (PKD2), accounting for approximately 85% and 15% of cases respectively12,13. Furthermore, the gene products for PKD1 and -2 localize to the cilium and are fundamental to ciliogenesis14,15. There is now a recognized family of human genetic disorders, known as the ciliopathies, which affect cilia function and result in CKD16.
The growing number of human genetic diseases affecting ciliary development and function is drawing global interest in this once considered vestigial organelle. The cilium, a hair-like cellular protrusion, is enriched with receptors and ion channels necessary for the transduction of key cell signaling events. The cilium consists of a microtubule-based axoneme, typically structured into nine radially arranged microtubule doublets with or without a central pair of singlet microtubules. The axonemal structure defines the type and mode of ciliary action. The 9+2 microtubule arrangement confers motility to the cilium where it is utilised in the movement of fluids across epithelial surfaces. The 9+0 configuration is non motile but is believed to mainly function in cellular signalling events17. Apart from CKD, the consequences of ciliary dysfunction are a set of characteristic ciliopathy features that include, obesity, retinal degeneration, polydactyly, and cognitive impairment16. However, CKD is amongst the most detrimental to the patient&#8217;s quality of life and therefore a major driving force behind the development of appropriate in vivo models for ciliary related CKD.
The zebrafish is an excellent model to understand the etiology of human genetic disease. Their quick development, production of large number of eggs, transparent tissue, and ex utero growth allows zebrafish developmental processes to be visualized and biological events manipulated with considerable ease. Genes can be genetically altered using the recent success of genome editing tools (CRISPR18 and TALENS19), knocked down using antisense morpholino technology20, or pharmacologically regulated by the addition of compounds to their aquatic environment. Indeed, zebrafish offer a platform to undertake experiments that are not permissive in other animal models. Whilst zebrafish are relatively simple vertebrates (compared to humans) they share many functionally conserved organs, genes, and signaling processes in common with humans. For example, the zebrafish kidney is remarkably similar in structure and function compared to humans21,22. However, unlike the mammalian kidney that develops through a succession of phases, each marked by a more developed kidney (pronephros, mesonephros, and metanephros), the embryonic zebrafish only develops a pronephros, the most immature form of a kidney. Whilst millions of nephrons can be found forming the building blocks of the mammalian kidney, the zebrafish embryo only possess two. The glomeruli, which allow for the initial blood filtrate, are fused at the midline just ventral to the aorta. Blood filters through the glomeruli into the pronephric tubules that run caudally along the axis, fusing prior to exit via the cloaca. The pronephric tubules are heavily ciliated with motile cilia that are permissive to the flow of filtrate towards the caudal exit3,4. This simple pronephric structure maintains zebrafish homeostasis through several weeks of larval growth where they eventually develop into a more complex mesonephros structure21. However, the zebrafish never develops a metanephros21. Despite the zebrafish idiosyncrasies, the zebrafish nephron is segmented with gene expression profiles equal to that observed in mammals and thus offers an unrivalled in vivo model for nephrogenesis3,22.
Routinely patients are tested for kidney function through a series of blood and urinary tests. Typically the blood is analyzed for dissolved salts, urea and creatinine. High levels of urea, creatinine and abnormal salt concentrations are indicative of problems with kidney function. Urinalysis using a colorimetric dipstick detects abnormal levels of protein, blood, pus, bacteria and sugar present in urine samples. Such tests normally require approximately 30 ml of urine or 5 - 10 ml of blood. It has been difficult to translate these types of assays to small in vivo model organisms, such as the zebrafish, mainly due to the impossible nature of collecting sufficient blood or urine to perform the assay. Here, we address the lack of appropriate zebrafish kidney function tests and describe an innovative technique for its study. By injecting a fluorescent dye into the blood stream we are able to monitor and individually quantify over time the filtration and excretion of fluorescent activity from the blood via the kidney. This method can be used to study kidney damage caused by disease, which we provide an example of.

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			<td height="21" style="height:21px;width:269px;">Name of Material/ Equipment</td>
			<td style="width:136px;">Company</td>
			<td style="width:136px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
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			<td height="42" style="height:42px;width:269px;">P-97 SUTTER Flaming/Brown type micropipette puller</td>
			<td style="width:136px;">Intracel</td>
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			<td height="21" style="height:21px;">Dumont #5 forceps&#160;</td>
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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.

Bardet-Biedl syndrome (BBS) is a rare heterogeneous ciliopathy that affects approximately 1:160,000 people worldwide16. Patients present with a number of associated problems including polycystic kidneys, subsequently patients frequently require kidney dialysis or transplantation24. ESRF is the most common cause of death in BBS, with around 30% of patients developing CKD16. Currently, 20 non-related genes have been implicated in BBS with no published genotype-phenotype association. The BBS...

Watch this Scientific Journal Video about Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay at JoVE.com

Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay

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 ...

evaluation-zebrafish-kidney-function-using-fluorescent-clearance

Research

JoVE Journal

Biology

13.1K Aufrufe.  University College London.  Der Zebrafisch ist ein beliebtes Tool zur chronischen Nierenerkrankung (CKD) zu modellieren. Allerdings macht ihre geringe Größe es unmöglich, die Nierenfunktion zu bewerten mit traditionellen Methoden. Wir beschreiben einen Fluoreszenzfarbstoff Nieren-Clearance-Test 1, die quantitative Analyse von Zebrafisch Nierenfunktion bei CKD ermöglicht. 

Serie: Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay

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 ...

Microinjection of Zebrafish Embryos to Analyze Gene Function

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic oligonucleotides with antisense complementarity to target RNAs, can be added to the embryo to reduce the expression of a particular gene product.  Conversely, processed mRNA can be added to the embryo to increase levels of a gene product.  The vehicle for adding either mRNA or morpholino to an embryo is microinjection.  Microinjection is efficient and rapid, allowing for the injection of hundreds of embryos per hour.  This video shows all the steps involved in microinjection.  Briefly, eggs are collected immediately after being laid and lined up against a microscope slide in a Petri dish.  Next, a fine-tipped needle loaded with injection material is connected to a microinjector and an air source, and the microinjector controls are adjusted to produce a desirable injection volume.  Finally, the needle is plunged into the embryo's yolk and the morpholino or mRNA is expelled.

This video shows how morpholino or mRNA can be injected into zebrafish embryos at the one-cell stage to decrease or increase the level of specific gene products during subsequent development.

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic ...

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. The protocol is a modified version of the standard in situ hybridization using alkaline phosphatase and substrates such as NBT/BCIP and Fast Red 1,2. This protocol utilizes standard digoxygenin and fluorescein labeled probes along with tyramide signal amplification (TSA) 3. The commercially available TSA kits allow flexible experimental design as fluorescence emission from green to far-red can be used in combination with various nuclear stains, such as propidium iodide, or fluorescence immunohistochemistry for proteins. TSA produces a reactive fluorescent substrate that quickly covalently binds to moieties, typically tyrosine residues, in the immediate vicinity of the labeled antisense riboprobe. The resulting staining patterns are high resolution in that subcellular localization of the mRNA can be observed using laser scanning confocal microscopy 3,4. One can observe nascent transcripts at the chromosomal loci, distinguish nuclear and cytoplasmic staining and visualize other patterns such as cortical localization of mRNA. Studies in Drosophila indicate that roughly 70% of mRNAs exhibit specific patterns of subcellular localization that frequently correlate with the function of the encoded protein 5. When combined with computer-aided reconstruction of 3D confocal datasets, our protocol allows the detailed analysis of mRNA distribution with sub-cellular resolution in whole vertebrate embryos.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. We include a propidium iodide nuclear counter-stain to highlight tissue organization.

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology.  Here, we present a high-resolution double ...

Using the optokinetic response to study visual function of zebrafish

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the visual functions of larval zebrafish1-5. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adult zabrafish. Here, we introduce how to measure the OKR of adult zebrafish with our simple custom-built apparatus using a new protocol which is established in our lab. Both our apparatus and step-by-step procedure of OKR in adult zebrafish are illustrated in this video. In addition, the measurements of the larval OKR, as well as the optomotor response (OMR) test of adult zebrafish, are also demonstrated in this video. This OKR assay of adult zebrafish in our experiment may last for up to 4 hours. Such OKR test applied in adult fish will benefit to visual function investigation more efficiently when the adult fish vision system is manipulated.
Su-Qi Zou and Wu Yin contributed equally to this paper.

Optokinetic response has been widely used to assess the visual functions of larval zebrafish. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adults1-5. Here, we introduce how to measure the OKR of adult zebrafish using a new protocol which is established in our lab.

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the ...

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 ...

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 Mammary Gland Development and Function in Mouse Models

The human mammary gland is composed of 15-20 lobes that secrete milk into a branching duct system opening at the nipple. Those lobes are themselves composed of a number of terminal duct lobular units made of secretory alveoli and converging ducts1. In mice, a similar architecture is observed at pregnancy in which ducts and alveoli are interspersed within the connective tissue stroma. The mouse mammary gland epithelium is a tree like system of ducts composed of two layers of cells, an inner layer of luminal cells surrounded by an outer layer of myoepithelial cells denoted by the confines of a basement membrane2. At birth, only a rudimental ductal tree is present, composed of a primary duct and 15-20 branches. Branch elongation and amplification start at the beginning of puberty, around 4 weeks old, under the influence of hormones3,4,5. At 10 weeks, most of the stroma is invaded by a complex system of ducts that will undergo cycles of branching and regression in each estrous cycle until pregnancy2. At the onset of pregnancy, a second phase of development begins, with the proliferation and differentiation of the epithelium to form grape-shaped milk secretory structures called alveoli6,7. Following parturition and throughout lactation, milk is produced by luminal secretory cells and stored within the lumen of alveoli. Oxytocin release, stimulated by a neural reflex induced by suckling of pups, induces synchronized contractions of the myoepithelial cells around the alveoli and along the ducts, allowing milk to be transported through the ducts to the nipple where it becomes available to the pups 8. Mammary gland development, differentiation and function are tightly orchestrated and require, not only interactions between the stroma and the epithelium, but also between myoepithelial and luminal cells within the epithelium9,10,11. Thereby, mutations in many genes implicated in these interactions may impair either ductal elongation during puberty or alveoli formation during early pregnancy, differentiation during late pregnancy and secretory activation leading to lactation12,13. In this article, we describe how to dissect mouse mammary glands and assess their development using whole mounts. We also demonstrate how to evaluate myoepithelial contractions and milk ejection using an ex-vivo oxytocin-based functional assay. The effect of a gene mutation on mammary gland development and function can thus be determined in situ by performing these two techniques in mutant and wild-type control mice.

This method describes how to dissect and assess mammary gland development and function from mice. Excised mammary glands are assessed for the degree of development using whole mount while milk ejection is evaluated using an oxytocin-based myoepithelial cell contraction assay.

The human mammary gland is composed of 15-20 lobes that secrete milk into a branching duct system opening at the nipple. Those lobes are themselves ...

Dissection of the Adult Zebrafish Kidney

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem cells during normal homeostasis and disease states. The understanding of adult stem cells has broad reaching implications for the future of regenerative medicine1. For example, better knowledge about adult stem cell biology can facilitate the design of therapeutic strategies in which organs are triggered to heal themselves or even the creation of methods for growing organs in vitro that can be transplanted into humans1. The zebrafish has become a powerful animal model for the study of vertebrate cell biology2. There has been extensive documentation and analysis of embryonic development in the zebrafish3. Only recently have scientists sought to document adult anatomy and surgical dissection techniques4, as there has been a progressive movement within the zebrafish community to broaden the applications of this research organism to adult studies. For example, there are expanding interests in using zebrafish to investigate the biology of adult stem cell populations and make sophisticated adult models of diseases such as cancer5. Historically, isolation of the zebrafish adult kidney has been instrumental for studying hematopoiesis, as the kidney is the anatomical location of blood cell production in fish6,7. The kidney is composed of nephron functional units found in arborized arrangements, surrounded by hematopoietic tissue that is dispersed throughout the intervening spaces. The hematopoietic component consists of hematopoietic stem cells (HSCs) and their progeny that inhabit the kidney until they terminally differentiate8. In addition, it is now appreciated that a group of renal stem/progenitor cells (RPCs) also inhabit the zebrafish kidney organ and enable both kidney regeneration and growth, as observed in other fish species9-11. In light of this new discovery, the zebrafish kidney is one organ that houses the location of two exciting opportunities for adult stem cell biology studies. It is clear that many outstanding questions could be well served with this experimental system. To encourage expansion of this field, it is beneficial to document detailed methods of visualizing and then isolating the adult zebrafish kidney organ. This protocol details our procedure for dissection of the adult kidney from both unfixed and fixed animals.
Dissection of the kidney organ can be used to isolate and characterize hematopoietic and renal stem cells and their offspring using established techniques such as histology, fluorescence activated cell sorting (FACS)11,12, expression profiling13,14, and transplantation11,15. 
We hope that dissemination of this protocol will provide researchers with the knowledge to implement broader use of zebrafish studies that ultimately can be translated for human application.

The zebrafish kidney is home to both renal and hematopoietic adult stem/progenitor cells, and represents an outstanding opportunity to study these cell types and their progeny in a vertebrate model organism. Here, we demonstrate a detailed dissection procedure that enables the researcher to identify and surgically remove the adult zebrafish kidney, which can be used for applications such as cell isolation, transplantation, and expression studies of kidney and/or blood cell populations.

Researchers working in the burgeoning field of adult stem cell biology seek to understand the signals that regulate the behavior and function of stem ...

A Fluorescent Screening Assay for Identifying Modulators of GIRK Channels

G protein-gated inward rectifier K+ (GIRK) channels function as cellular mediators of a wide range of hormones and neurotransmitters and are expressed in the brain, heart, skeletal muscle and endocrine tissue1,2. GIRK channels become activated following the binding of ligands (neurotransmitters, hormones, drugs, etc.) to their plasma membrane-bound, G protein-coupled receptors (GPCRs). This binding causes the stimulation of G proteins (Gi and Go) which subsequently bind to and activate the GIRK channel. Once opened the GIRK channel allows the movement of K+ out of the cell causing the resting membrane potential to become more negative. As a consequence, GIRK channel activation in neurons decreases spontaneous action potential formation and inhibits the release of excitatory neurotransmitters. In the heart, activation of the GIRK channel inhibits pacemaker activity thereby slowing the heart rate.
GIRK channels represent novel targets for the development of new therapeutic agents for the treatment neuropathic pain, drug addiction, cardiac arrhythmias and other disorders3. However, the pharmacology of these channels remains largely unexplored. Although a number of drugs including anti-arrhythmic agents, antipsychotic drugs and antidepressants block the GIRK channel, this inhibition is not selective and occurs at relatively high drug concentrations3.
Here, we describe a real-time screening assay for identifying new modulators of GIRK channels. In this assay, neuronal AtT20 cells, expressing GIRK channels, are loaded with membrane potential-sensitive fluorescent dyes such as bis-(1,3-dibutylbarbituric acid) trimethine oxonol [DiBAC4(3)] or HLB 021-152 (Figure 1). The dye molecules become strongly fluorescent following uptake into the cells (Figure 1). Treatment of the cells with GPCR ligands stimulates the GIRK channels to open. The resulting K+ efflux out of the cell causes the membrane potential to become more negative and the fluorescent signal to decrease (Figure 1). Thus, drugs that modulate K+ efflux through the GIRK channel can be assayed using a fluorescent plate reader. Unlike other ion channel screening assays, such atomic absorption spectrometry4 or radiotracer analysis5, the GIRK channel fluorescent assay provides a fast, real-time and inexpensive screening procedure.

A real-time screening procedure for identifying drugs that interact with G protein-gated inward rectifier K+ (GIRK) channels is described. The assay utilizes membrane potential-sensitive fluorescent dyes to measure GIRK channel activity. This technique is adaptable for use on a number of cell lines.

G protein-gated inward rectifier K+ (GIRK) channels function as cellular mediators of a wide range of hormones and neurotransmitters and are expressed in ...

Analysis of Nephron Composition and Function in the Adult Zebrafish Kidney

The zebrafish model has emerged as a relevant system to study kidney development, regeneration and disease. Both the embryonic and adult zebrafish kidneys are composed of functional units known as nephrons, which are highly conserved with other vertebrates, including mammals. Research in zebrafish has recently demonstrated that two distinctive phenomena transpire after adult nephrons incur damage: first, there is robust regeneration within existing nephrons that replaces the destroyed tubule epithelial cells; second, entirely new nephrons are produced from renal progenitors in a process known as neonephrogenesis. In contrast, humans and other mammals seem to have only a limited ability for nephron epithelial regeneration. To date, the mechanisms responsible for these kidney regeneration phenomena remain poorly understood. Since adult zebrafish kidneys undergo both nephron epithelial regeneration and neonephrogenesis, they provide an outstanding experimental paradigm to study these events. Further, there is a wide range of genetic and pharmacological tools available in the zebrafish model that can be used to delineate the cellular and molecular mechanisms that regulate renal regeneration. One essential aspect of such research is the evaluation of nephron structure and function. This protocol describes a set of labeling techniques that can be used to gauge renal composition and test nephron functionality in the adult zebrafish kidney. Thus, these methods are widely applicable to the future phenotypic characterization of adult zebrafish kidney injury paradigms, which include but are not limited to, nephrotoxicant exposure regimes or genetic methods of targeted cell death such as the nitroreductase mediated cell ablation technique. Further, these methods could be used to study genetic perturbations in adult kidney formation and could also be applied to assess renal status during chronic disease modeling.

The zebrafish adult kidney is an excellent system for renal regeneration and disease studies. An essential aspect of such research is the assessment of nephron structure and function. This protocol describes several methodologies that can be implemented to assess nephron tubule composition and to evaluate renal reabsorption.

The zebrafish model has emerged as a relevant system to study kidney development, regeneration and disease. Both the embryonic and adult zebrafish kidneys ...