In vivo analysis of renal epithelial cells in zebrafish

CHAPTER

In vivo analysis of renal epithelial cells in zebrafish

10

Yuanyuan Li, Wenyan Xu, Stephanie Jerman, Zhaoxia Sun* Department of Genetics, Yale University School of Medicine, New Haven, CT, United states *Corresponding author: e-mail address: [email protected]

Chapter outline 1 Introduction......................................................................................................164 2 Cell biology of renal epithelial cells in the zebrafish embryo................................165 2.1 Lumen formation and the establishment of apical-basal polarity..............165 2.2 Ciliogenesis.......................................................................................166 2.3 Planar cell polarity.............................................................................167 2.4 Morphogenic movement......................................................................167 3 Methods to analyze renal epithelial cells in the zebrafish pronephros...................168 3.1 CRISPR/Cas9-mediated gene inactivation.............................................168 3.1.1 Materials........................................................................................169 3.1.2 Protocol..........................................................................................170 3.2 Evaluate genetic variants with rescue experiment in zebrafish.................173 3.2.1 Materials........................................................................................174 3.2.2 Protocol..........................................................................................174 3.3 Evaluate pronephric epithelial cells......................................................175 3.3.1 Method 1. Immunostaining and visualization of whole-mount embryos...176 3.3.2 Method 2. Cryosection and immunostaining to visualize structures of kidney epithelial cells..........................................................................178 4 Discussion and future direction..........................................................................179 Acknowledgments..................................................................................................179 References............................................................................................................179

Abstract The zebrafish kidney has been used effectively for studying kidney development, repair and disease. New gene editing capability makes it a more versatile in vivo vertebrate model system to investigate renal epithelial cells in their native environment. In this chapter we focus on dissecting gene function in basic cellular biology of renal epithelial cells, including lumen formation and cell polarity, in intact zebrafish embryos. Methods in Cell Biology, Volume 154, ISSN 0091-679X, https://doi.org/10.1016/bs.mcb.2019.04.016 © 2019 Elsevier Inc. All rights reserved.

163

164

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

1 Introduction The highly polarized epithelial cell is a main component of the vertebrate kidney, a vital organ for removing waste, maintaining water and electrolyte homeostasis, and modulating blood pressure. Lining distinct renal segments, epithelial cells not only provide structural integrity of the fluid discharge system, they also function actively in secretion, reabsorption and tissue repair. Moreover, through cell-cell interaction, paracrine and endocrine regulation, and the sympathetic and parasympathetic nervous system, kidney epithelial cells are an integral part of a complex network that allows the organism to adapt dynamically to the changing environment and functional requirements. Cultured epithelial cells have been invaluable in teasing out the basic cell biology, including the proliferation, differentiation, apical-basal polarity and ciliogenesis, of renal epithelial cells. In a complementary fashion, in vivo models allow investigation of renal epithelial cells in their native environment and can address interconnectivity of cells at a macroscopical level. They have been used successfully to investigate the mechanisms underlying the planar cell polarity (PCP), morphogenic movement and mesenchymal-epithelial interaction during kidney development, homeostasis, damage repair and regeneration (Carroll & Das, 2011; Carroll & Yu, 2012; Chang-Panesso & Humphreys, 2017; Combes, Davies, & Little, 2015; Das et al., 2013; Karner et al., 2009; Kramann et al., 2017; Lienkamp et al., 2012). The zebrafish Danio rerio is an established vertebrate model organism for studying kidney development and disease (Drummond, 2002; Drummond & Davidson, 2010; Jerman & Sun, 2017; Wingert & Davidson, 2008). The functional kidney in the zebrafish embryo is the pronephros. It contains a pair of nephrons with two fused glomeruli located ventral to the notochord and medial to the pair of pectoral fins. Although very simplified, the pronephros contains well differentiated podocytes and epithelial cells, which express different segment markers. The conservation of the basic components and function of the kidney, combined with the high fecundity and small size of zebrafish, has facilitated high throughput genetic and chemical screens for factors affecting kidney epithelial cells (Drummond et al., 1998; Sun et al., 2004). In recent years, rapid technical advancements in next-generation sequencing, human genetics, gene editing, organoid culture and live imaging have brought new opportunities and challenges. We posit that in this rapidly changing landscape the zebrafish remains a highly relevant, and arguably more powerful, model organism for renal studies. Most significantly, new gene editing capability makes it highly efficient to assess gene function and the functional consequence of genetic variants in an intact vertebrate animal. We refer readers to previous reviews for details of the development and structure of the zebrafish kidney (Drummond, 2002; Drummond & Davidson, 2010; Jerman & Sun, 2017; Wingert & Davidson, 2008). Here we introduce several key in vivo cell biology readouts of kidney epithelial cells in the zebrafish embryo and provide several protocols for dissecting gene function in this context.

2 Cell biology of renal epithelial cells in the zebrafish embryo

2 Cell biology of renal epithelial cells in the zebrafish embryo 2.1 Lumen formation and the establishment of apical-basal polarity The tubule and duct of the zebrafish pronephros form in situ from the intermediate mesoderm during development through mesenchyme to epithelium transition. During early somitogenesis stages, a number of transcription factors, including Lim1, Pax2 and vHnf1, are already expressed in specific domains of the intermediate mesoderm and specify future segments of the nephron (Serluca & Fishman, 2001; Sun & Hopkins, 2001). However, on the cellular level, the intermediate mesoderm at this stage appears as a cluster of cells lateral to the somites on histological sections stained with hematoxylin and eosin (HE). No lumen or polarity can be detected (Fig. 1A). At around the 16–17 somite stage, the forming tubule resembles a solid rod on most histological cross sections. However, apical-basal (AP) polarity can already be detected by immunostaining. Specifically, Laminin is enriched on the outside of the rod, while Cadherin 17 (Cdh17) predominantly displays a punctate pattern, likely representing intracellular vesicles (Fig. 1B, upper panel). By the 24-somite stage, a central lumen is readily detectable (Fig. 1A). By 35 h post

FIG. 1 Formation of the pronephric tubule in zebrafish. (A). Mesenchyme to epithelium transition. Upper panel is a diagram and the lower panel shows HE cross sections. At the 6-somite (6S) stage, the intermediate mesoderm (IM, lined with dotted line) is lateral to the somite (SM). Half of the neural keel (NK) is also visible on this section. At the 16-somite (16S) stage, a solid rod without lumen is seen. At the 24-somite (24S) stage, a lumen (red arrow) is obvious at the center. (B) Lateral view of the pronephric tubule. The upper panel shows a tubule at the 18-somite stage (18S) stained with anti-Lamin (Lam) in red and Chd17 in green. The lower panel shows a tubule at 35 hpf stained with aPKC in red and Cdh17 in green. (C) Cross sections of a tubule at 50 hpf. The upper panel shows filamentous actin (Fact) labeled by phalloidin in red and Chd17 in green. The lower panel shows aPKC in red and Na+/K+ ATPase labeled with the α6F antibody in green. Panel (C) is reproduced from fig. 1I in Duldulao, N. A., Lee, S., & Sun, Z. (2009). Cilia localization is essential for in vivo functions of the Joubert syndrome protein Arl13b/scorpion. Development, 136(23), 4033–4042.

165

166

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

fertilization (hpf ), Cdh17 is enriched on the basal-lateral membrane of the epithelial cells, while atypical PKC is enriched at the apical side (Fig. 1B, lower panel). By 40–48 hpf , the entire tubule and duct is patent and glomerular filtration is established (Drummond et al., 1998). The apical-basal polarity can also be detected by Na+/K+ ATPase at the basal lateral side and filamentous actin at the apical side (Fig. 1C).

2.2 Ciliogenesis Cilia on zebrafish epithelial cells are motile and the pronephric tube contains both multi and single ciliated cells (MCCs and SCCs), a significant difference from the mammalian metanephros. Ciliogenesis in zebrafish pronephric epithelial cells occur in two waves. In the first wave, which occurs almost simultaneously with lumen formation, solitary cilia emerge from newly differentiated epithelial cells (Fig. 2A). Starting from 24 hpf, although most cells are single ciliated, clusters of basal bodies in individual cells in the proximal tubule can be detected, indicating that multi-ciliogenesis in the second wave has begun. By 30 hpf, multi-cilia can be

FIG. 2 Ciliogenesis in the pronephric tubule. (A) Cilia (anti-Arl13b in green) and basal bodies label by anti-γtubulin (γtub) in red. At the 24-somite (24S) stage, most are single ciliated. Clusters of basal bodies (yellow arrow heads) can be seen at 24 hpf and 30 hfp. (B) Cilia bundle labeled with anti-acetylated tubulin (a-tub) in red in a 5 dpf pronephric tubule labeled with anti-Cdh17 in green. (C) A wild type (WT) and arl13bhi459 mutant (MUT) stained with anti-acetylated tubulin and detected with DAB reaction. Arrow points to cilia bundle in the pronephric tubule. Panel (A) is reproduced from fig. 5B–D in Li, J., & Sun, Z. (2011). Qilin is essential for cilia assembly and normal kidney development in zebrafish. PLoS One, 6(11), e27365. Panel (B) is reproduced from fig. 4E in Duldulao, N. A., Lee, S., & Sun, Z. (2009). Cilia localization is essential for in vivo functions of the Joubert syndrome protein Arl13b/scorpion. Development, 136(23), 4033–4042. Panel (C) is reproduced from fig. S2A, B in Duldulao, N. A., Lee, S., & Sun, Z. (2009). Cilia localization is essential for in vivo functions of the Joubert syndrome protein Arl13b/scorpion. Development, 136(23), 4033–4042.

2 Cell biology of renal epithelial cells in the zebrafish embryo

detected in this region (Fig. 2A). In the mature tubule, cilia form bundles in the lumen (Fig. 2A), which beat coordinately to efficiently discharge fluid from this aquatic animal (Kramer-Zucker et al., 2005). The cells in the most posterior region, corresponding to the collecting duct, remain single ciliated. Mutations in genes essential for cilia biogenesis could show differential impact on the two waves of ciliogenesis in the zebrafish pronephros, mainly caused by maternal contribution of gene products in zebrafish oocytes. As a consequence, zygotic homozygous mutant offspring from a heterozygous mother will start with wild type gene products. Because of the rapid development of the zebrafish embryo, maternal contribution may mask early functional requirements for gene function before the decay of the deposited mRNA and protein. In the case of ciliogenesis in pronephric epithelial cells, maternal contribution may be able to support the early biogenesis of single cilia, but usually is insufficient for the second phase of multi-ciliogenesis, which occurs later and requires more building materials (Li & Sun, 2011). It is critical to analyze ciliogenesis at both stages to ascertain whether the gene of interest is required for cilia biogenesis.

2.3 Planar cell polarity In addition to apical-basal polarity, epithelial cells also show PCP, which refers to polarity within the epithelial sheet. This phenomenon is best illustrated by the proximal to distal orientation of hair bristles on the Drosophila wing, which has facilitated the dissection of the signaling pathway that establishes and maintains this polarity across a field of cells. As the PCP pathway share common components with the Wnt-β-catenin pathway, it is also known as the non-canonical Wnt pathway. The planar polarity of the zebrafish renal epithelial cells is detectable by the orientation of cilia pointing from the proximal to distal end, the direction of the flow. PCP is also critical for mammalian kidney development. It is required for proper cell division axis and tubule morphogenesis and defective PCP may contribute to cyst formation in polycystic kidney disease (Carroll & Das, 2011; Carroll & Yu, 2012; Karner et al., 2009; Lienkamp et al., 2012; Verdeguer et al., 2010).

2.4 Morphogenic movement Since the pair of pronephros form from two stripes of intermediate mesoderm, no branch morphogenesis is involved in the development of the zebrafish pronephros during the embryonic stage. Nonetheless, the developing pronephric tubule undergoes dynamic morphogenic movement. Live imaging showed that epithelia cells in the pronephric tubule migrates collectives toward the proximal side (Vasilyev et al., 2009). As a result, the proximal tubule becomes increasingly convoluted as the nephron matures (Fig. 3A). The convolution of the proximal tubule therefore can be used as a simple readout for directional morphogenic movement of pronephric epithelial cells. If a defect is detected, live imaging can be used to monitor this movement directly.

167

168

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

FIG. 3 Morphogenesis of the pronephric tubule. (A) Whole zebrafish embryos labeled with α6F (anti-Na+/K+ ATPase) and DAB reaction. At 57 hpf, the neck region (arrow) is visible. At 5 dpf, the proximal tubule (arrow) becomes very convoluted. (B) A zebrafish larva at 6 weeks (6wk) labeled with α6F (anti-Na+/K+ ATPase) and DAB reaction. Arrow points to convoluted tubule in the mesonephros. (C) A wild type (WT) and arl13bhi459 mutant (MUT) at 50 hpf stained with α6F (anti-Na+/K+ ATPase) and detected with DAB reaction. The tubule in the mutant is enlarged. Panel (C) is reproduced from fig. 1G and H in Duldulao, N. A., Lee, S., & Sun, Z. (2009). Cilia localization is essential for in vivo functions of the Joubert syndrome protein Arl13b/scorpion. Development, 136(23), 4033–4042.

3 Methods to analyze renal epithelial cells in the zebrafish pronephros 3.1 CRISPR/Cas9-mediated gene inactivation For the lack of ES cells, generating targeted mutations through homologous recombination was not feasible in zebrafish. Engineered zinc finger nuclease (ZFN) technique uses engineered zinc fingers to direct the FokI endonuclease to generate breaks in target sequences. Repair of DNA breaks is error prone and frequently leads to micro insertions and deletions (indels). However, the construction of zinc finger nuclease is not trivial and limits its use in zebrafish especially as a high throughput method (Meng et al., 2008). The programmable CRISPR system changed the picture completely (Hwang et al., 2013). Moreover, since zebrafish eggs are fertilized externally, microinjection into zebrafish eggs is technically straightforward. Because of cytoplasmic streaming, injected materials will be distributed ubiquitously in the rapidly developing embryo. Notably, sgRNA can be synthesized via in vitro transcription using annealed oligos and injected together with Cas9 mRNA or protein, completely eliminating the need of cloning for any targeted gene. The sole target specific reagent required is an oligo around 52 bp in length (Gagnon et al., 2014; Jao, Wente, & Chen, 2013). These features make CRISPR-Cas9 mediated gene inactivation in zebrafish efficient and cost effective. As an example, we co-injected two gRNAs against zmynd10 with Cas9 protein, and readily detected body curvature and kidney cysts, two phenotypes closely associated with cilia motility defects in zebrafish, in injected embryos (Fig. 4B and C).

3 Methods to analyze renal epithelial cells in the zebrafish pronephros

FIG. 4 CRISPR-Cas9 mutagenesis in zebrafish. (A) Upper panel shows PCR product used as template for in vitro transcription of sgRNA. Lower panel shows sgRNA product. Lane 1 is size marker. (B) Un-injected sibling control and (C) embryos injected with Cas9 and sgRNAs against zmynd10. (D) A sequencing result using whole embryo lysate. Target sequence is underlined.

On the DNA level, direct sequencing of PCR products using whole embryo lysates revealed overlapping peaks appearing abruptly in the target region, suggesting that insertions and deletions (indels) occur at very high frequency in injected F0 embryos (Fig. 4D). A number of studies have been carried out to optimize protocols for CRISPRCas9 mediated gene knockout in zebrafish (Burger et al., 2016; Gagnon et al., 2014; Jao et al., 2013; Shah et al., 2015). It is worth noting that CRISPR-Cpf1 has also been shown to work effectively in zebrafish and provides additional targets (Moreno-Mateos et al., 2017). The following describes the CRISPR/Cas9 method we use in our laboratory that incorporates various aspects of previous protocols and our own experience. In this method, commercial Cas9 protein is co-injected with sgRNA transcribed in vitro. Alternatively, mRNA encoding Cas9 can be used in place of Cas9 protein and crRNA/tracrRNA can be ordered directly from commercial sources.

3.1.1 Materials • • • •

Wild type adult zebrafish, both male and female Standard microinjection reagents and equipment as described (Yuan and Sun, 2009) Dissecting microscope Thermocycler

169

170

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

• • • • • • • • • • • • •

Standard DNA electrophoresis apparatus Qiagen gel purification kit Nanodrop Dumont forceps Cas9 protein with nuclear localization sequence (PNA-Bio CP01) GoTaq Green Master Mix (Promega M7122) HiScribe T7 Kit (NEB 2040S) Mini quick spin columns for RNA (Roche 1-814-427) 50 mM NaOH DNAse I 1 M Tris-HCl, not pH adjusted Phenol red (Sigma-Aldrich Co., P0290) 200 μL tubes

3.1.2 Protocol (1) Target selection. Use a sequence alignment program, such as MegAlign in DNASTAR or HomoloGene under NCBI, to identify conserved regions in the protein encoded by gene of interest. Select three to six targets by adding the CRISPRscan track on the UCSC Genome Browser, which allows for visualization of targets in the context of gene structures. The CRISPRscan platform is used because its scoring algorithm is built on results from similar experimental settings in zebrafish (Moreno-Mateos et al., 2015). Targets in each gene will be selected based on whether they are located in essential regions, conserved regions and regions more 50 side in the gene. (2) Verify that targeted regions can be readily genotyped. Some target regions are refractory to PCR because of redundant sequence or high GC content. Since multiple targets are generally available for each gene, we recommend testing whether the target region can be easily PCR amplified and sequenced before attempting gene editing. • Design primer pairs that flank each selected target site at a 100–400 bp distance. • Obtaining zebrafish genomic DNA by pooling five embryos in 50 μL of 50 mM NaOH. Place on a heat block at 95 °C for 10 min to lyse embryos. Neutralize with 10 μL 1 M Tris-HCl. Mix well and spin briefly. The lysed sample can now be stored at 20 °C for later use. • Take 1 μL lysate, set up 20 μL PCR reaction with 2XGoTaq green mix using a standard PCR protocol. • Run PCR samples on a 1.5% agarose gel to ensure a single robust band at the predicted size is present. • Use Qiagen gel purification kit to purify DNA, send for Sanger sequencing and verify that clean sequence result can be obtained. (3) Generate sgRNA via in vitro transcription. • Order the universal scaffolding primer 50 -ttttgcaccgactcggtgccactttttcaa gtTgataaCggactagccttattttaactt

3 Methods to analyze renal epithelial cells in the zebrafish pronephros

gctatttctagctctaaaac-30 and the gene-specific primer using the exact sequence shown in “Promoter + sgRNA + universal primer sequence” in CRISPscan. • Generate template for in vitro transcription via PCR. • Mix 5 μL GoTaq Green Master Mix, 2.5 μL 10 μm gene specific primer and universal scaffold primer each on ice. Run a standard PCR protocol with an annealing temperature of 45 °C. • Run 1 μL PCR product on a 2% agarose gel to verify the presence of a single robust band at around 120 bp. An example is shown in the upper lane of Fig. 4A. PCR products can be stored at 20°C and used later. • Synthesize sgRNA via in vitro transcription. As the PCR products contain the T7 promoter, they are used directly as template for sgRNA production via in vitro transcription using the HiScribe T7 kit from NEB. We routinely mix three templates for a single gene in a single in vitro transcription reaction when the yield of PCR product is relatively equivalent for each target. Below is the protocol using three pooled templates. Alternatively, this recipe can be adjusted for using a single template in each reaction. • Mix equal amount of ATP, GTP UTP and CTP (100 mM each) in the HiScribe T7 Kit to make NTP mix. • Set up 15 μL reaction by mix 6 μL NTP mix, 6 μL (2 μL of each template) PCR product, 1.5 μL 10 buffer, 1.5 μL enzyme mix. • Incubate at 37 °C for 2 h. • Add 1 μL DNAse I. Incubate at 37 °C for 30 min. • Add 10 μL H2O to reaction mix to make the total volume 25 μL. • Purify mRNA using mini quick spin column for RNA following the manufacturer’s protocol. • Evaluate sgRNA products. Although the product is RNA, we find a quick run on a routine DNA gel suffice to monitor the integrity of the product. To minimize potential RNA degradation, use fresh buffer, clean tray and a fast run. Specifically, run 2 μL of the product on a 1.5% agarose/TAE gel for 15 min at 120 V with 1  loading dye and nuclease-free water. The resulting band should migrate at approximately 100–120 bp. An example is shown in the lower panel of Fig. 4A. • Use a Nanodrop to determine sgRNA concentration under the RNA setting. • Aliquot, label with concentration and date. Store at 80 °C. (4) Microinjection of zebrafish embryos. A detailed microinjection protocol has been described before (Yuan and Sun, 2009). Here we provide guidelines specific for microinjection of CRISPR/Cas9 reagents. We routinely pool three sgRNAs against the same gene together in the initial test, although it has been shown that up to 12 sgRNAs can be pooled (Shah et al., 2015). • In the afternoon before the day of micro-injection, place pairs of adult zebrafish in mating boxes. Separate male and female fish with a divider. On the morning of the injection day, remove divider, observe and collect fertilized eggs according to standard protocols (Westerfield, 1993).

171

172

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

Prepare 4 μL injection mixture to a final concentration of 500 ng/μL Cas9 protein, 150 ng/μL sg RNA, 1XCas9 buffer, 300 mM KCl and 0.05% phenol red. Incubate mixture at RT for 5 min before moving onto ice. Load injection needle. • Perform micro-injection into zebrafish embryos before the four-cell, but preferably at the one-cell stage. Keep a portion of siblings from the same clutch as the un-injected control. • Observe injected embryos up to 5 dpf. • Genotype pooled embryos as described in Step 2. Indels will lead to scrambling of the sequence chromatogram. An example is shown in Fig. 4D. In contrast to the clean trace in the wild type sample, additional overlapping peaks appear in injected embryos. The webtool Tracking of Indels by Decomposition (TIDE) (https://tide.deskgen.com/) can then be used to deconvolute traces and estimate indel frequency (Brinkman et al., 2014). • Perform biological repeats. • If consistent and interesting phenotypes are observed, inspect sequencing result of each target to eliminate any sgRNA in the pool that is inefficient in introducing indels. Individual sgRNA that induces indels effectively can be injected to verify its potency to induce phenotypes. Moreover, the exact indel needs to be identified by cloning PCR product and sequencing individual clones. We refer readers to a previous protocol for this step (Jao et al., 2013). (5) Interpretation of positive results. Although the CRISPR-Cas9 system shows relatively low toxicity in zebrafish, non-specific effects are still possible. Sufficient controls need to be included to rule out toxicity and off-target effect especially when phenotypes are evaluated in injected F0 embryos. The following are a number of possible controls in ascending levels of stringency: (A) uninjected sibling control, (B) co-injection of sgRNA against an unrelated target with Cas9 and (C) sgRNA against different targets in the same gene. (6) Interpretation of negative results. Quite frequently, small indels, even when outof-frame and in conserved regions, lead to no obvious phenotypes. Similar to positive results, negative results need to be interpreted with caution. In addition to functional redundancy, alternative splicing, translational initiation and ribosome readthrough could lead to production of functional products and false negative results. (7) Generate and evaluate stable lines. A stable line of carriers not only provide a steady supply of materials, it can also serve as an effective tool to validate phenotypes observed in F0. Since zebrafish has 25 pairs of chromosomes, offtarget mutations can be segregated out in stable lines. To obtain viable carriers, inject suboptimal doses of sgRNA and Cas9 into wild type embryos. Lyse a subpool of the injected embryos at 3 dpf to extract genomic DNA. Perform PCR to amplify the target region and use Sanger sequencing to verify successful mutagenesis in the target region. Raise the rest of the injected embryos to •

3 Methods to analyze renal epithelial cells in the zebrafish pronephros

adulthood. Screen F1 fish for carriers of indel mutations. Cross positive carriers with the same indel to generate homozygous mutants and inspect mutants to verify that the phenotypes observed in F0 is reproduced.

3.2 Evaluate genetic variants with rescue experiment in zebrafish The rapid advancement of sequencing technology has revolutionized human genetic studies. Up to date, most responsible genes for common Mendelian genetic diseases in human have been identified. The challenge is how to evaluate functional contribution/involvement of genetic variants identified in rare or non-Mendelian diseases in a relatively high-throughput fashion. The zebrafish is a vertebrate system with conserved structures of the kidney and genes involved in kidney development and disease are usually well conserved in this organism. In addition, genetic mutants already exist for many of the genes or can be readily generated via CRISPR/Cas9 as described above. Furthermore, mRNA can be synthesized in vitro and injected into zebrafish eggs in adjustable dosages. Based on these factors, we reason that rescue experiment in zebrafish via mRNA microinjection can be used effectively to evaluate the function of genetic variants both qualitatively and quantitatively. Using this method, we were able to contribute to the identification of IFT27 as a Bardet-Biedl syndrome gene, CLUAP1 as a Leber congenital amaurosis gene and PIH1D3 as a primary ciliary dyskinesia gene (Aldahmesh et al., 2014; Olcese et al., 2017; Soens et al., 2016). The prerequisite for using rescue experiment to test variant function is that re-expression of the wild type protein using the same method can rescue a lossof-function (LOF) mutant. In our experience, proteins with structural roles generally can rescue mutants without causing deleterious consequences in the developing embryo when expressed ubiquitously via mRNA injection. In contrast, enzymes or other proteins with instructive functions in signaling pathways, such as β-catenin, more likely will disrupt normal development when expressed using the same method. This issue potentially can be addressed by dosing the amount of injected mRNA carefully. It is also possible to perform rescue experiment by tissue specific expression via transgene. However, the latter is more labor intensive and here we limit our scope to transient expression and rescue via mRNA injection. We frequently start with mRNA encoding untagged wild type zebrafish protein of interest. After verifying the feasibility of rescue experiment, adding an epitope tag such as eGFP will facilitate the monitoring of gene expression. The tagged version also needs to be validated through rescue experiment. It may be necessary to change the location of the tag to obtain a functional version. It is also possible to rescue zebrafish mutants with the re-expression of the human homolog, although in our experience the zebrafish version tends to be more potent in this context. If the tested genetic variation is localized in a conserved region, using mRNA encoding the zebrafish homolog and the corresponding variant is a logic choice. Below we describe the general protocol of rescue experiment using mRNA injection.

173

174

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

3.2.1 Materials • • • • • • • • • • • • • •

Standard microinjection reagents and equipment as described previously (Yuan & Sun, 2009). Heterozygous carriers of LOF mutation in the gene of interest mMESSAGE mMACHINE kit (Ambion, AM1340) Tris lysis buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 0.5% NP-40, 0.1 mM DTT) Protease inhibitor (Roche 11836170001) Sample buffer (2% SDS, 100 mM DTT and 2.5% β-mercaptoethanol) SDS-PAGE gel (Biorad or other vendors) Mini-Protean Gel system (Biorad) Anti-GFP antibody (Roche 11814460001) Anti-tubulin (Molecular Probes, A11126) Anti-IgG-horseradish peroxidase (HRP) conjugate (Jackson Immuno-Research) Standard Western blot apparatus Western Lightning Detection Kit (PerkinElmer Life Sciences) Restore stripping buffer (ThermoFisher)

3.2.2 Protocol (1) In vitro transcription of capped mRNA. We routinely sub-clone the coding sequence of the gene of interest into the pCS2 + Vector, which carries a SP6 promoter for in vitro transcription and a Poly(A) sequence that improves the stability and translational efficiency of the mRNA. Similar protocols as sgRNA synthesis described above will be used for in vitro transcription, RNA purification and quantification, with a few modifications. To be used as a template for in vitro transcription, plasmid DNA needs to be linearized via restriction digestion at a site after the Poly(A) sequence and purified either via Qiagen columns or phenol-chloroform extraction. In addition, capped mRNA will be used in this experiment. Accordingly, an in vitro transcription kit compatible with capping, such as the mMESSAGE mMACHINE kit, will be used. (2) Rescue experiment. Obtain fertilized zebrafish eggs from heterozygous carries through natural spawning. Perform microinjection as described before (Yuan & Sun, 2009). 250 pg total mRNA is a good starting point based on previous experience. Keep a portion of siblings as uninjected control. Raise embryos, observe whether eGFP signal can be detected under a fluorescent dissection scope between the shield to bud stage. Observe embryos up to 5 dpf. Count both phenotypic and non-phenotypic embryos. In addition, record whether the severity of phenotypes has changed. (3) Statistical analysis: 25% offspring from crosses between heterozygous carriers are expected to be homozygous mutants. Fisher’s exact test can be used to

3 Methods to analyze renal epithelial cells in the zebrafish pronephros

exam whether the experimental group is significantly different from the expected and the control group using free webpages such as https://www. langsrud.com/fisher.htm, or R packages. (4) Quantification of the activity of the tested variant. If a variant is able to rescue the mutants, it may be a non-pathogenic allele. Alternative, it could be a partial LOF or hypomorphic allele. A dosage assay will be able to distinguish these two possibilities. Inject a deceasing amount of wild type and variant mRNA until reaching the minimal dose that can rescue significantly and compare (Duldulao et al., 2009). (5) Quantify expression level via Western blot. Although eGFP signal can be used to conveniently monitor the expression of the tested protein in live embryos, this assay is not quantitative. Western blot can be used to quantify whether the tested variant lead to a changed level of protein. Deyolk and homogenize embryos between 6 and 24 hpf in Tris lysis buffer containing protease inhibitors as described previously (Link, Shevchenko, & Heisenberg, 2006). Denature proteins in SDS sample buffer at 100 °C for 7 min and clear by centrifugation at 12,000g for 3 min. Load a series of different amount onto SDS-PAGE gel. Perform electrophoresis and transfer to a PVDF membrane. Blot with anti-eGFP antibody, followed by HRP-conjugated secondary antibody and expose with the Western detection kit. Strip the blot using stripping buffer and re-probe with anti-tubulin. Quantify band intensity with ImageJ, normalize with tubulin signal.

3.3 Evaluate pronephric epithelial cells The simple pronephros in the zebrafish embryo is well suited for evaluating kidney epithelial cells in their native environment. A number of biomarkers have been developed for visualizing structures or components of the kidney epithelial cells. For example, GFP-tagged pleckstrin homology (PH) domain of phospholipase Cd1 (PH-Plcd1-GFP) and PH-Akt1-GFP has been used to visualize PtdIns(4,5)P2 and PtdIns(3,4,5)P3, respectively (Xu et al., 2017). Here we focus on immunostaining as it does not require microinjection, crossing of transgenic fish or live imaging and multiple samples can be processed simultaneously. Due to the small size of the embryo, whole embryos can be stained, flat-mounted and visualized. This method is suitable for obtaining a macroscopic view of the pronephros, including the size and morphology of the tubule and duct, and the status of cilia bundle formation (Figs. 2C and 3C). It also allows visualization of the apical-basal polarity of renal epithelial cells in a side view (Figs. 1B and 2B). By using a partially dissected 6-week-old zebrafish, we were able to visualize almost the whole epithelial tube, including the convoluted region in the pronephros and the mesonephros (Fig. 3B). Alternatively, sections can be used to visualize the lumen and AP polarity in a cross view. Below we provide protocols for both methods.

175

176

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

3.3.1 Method 1. Immunostaining and visualization of whole-mount embryos 3.3.1.1 Materials • • • • • • • • • • • • • • • • • • • •

Medical grade tricaine (Western Chemical, Inc) Formalin Dent’s fixative (80% methanol and 20% DMSO) 2 mL microcentrifuge tubes PBST (PBS and 0.05% Tween-20) blocking buffer (PBST with 10% normal serum) Methanol DMSO 30% H2O2 solution Acetone 0.05% trypsin Normal serum Primary antibodies (Table 1) Anti-IgG-HRP conjugate (Pierce) 3,30 -Diaminobenzidine tablet (D-5905, Sigma-Aldrich) BABB (two parts benzyl benzoate and one part benzyl alcohol) Alexa 488 and Alexa 594 conjugated secondary antibodies (Invitrogen) ProLong™ Gold Antifade Mountant with DAPI (ThermoFisher) Superfrost Plus microscope slides (Fisher Scientific) Glass coverslips (Fisher Scientific)

Table 1 Markers for renal epithelial cells in zebrafish. Name +

+

Na /K ATPase Laminin aPKC Cdh17 F-Actin Acetylated tubulin Arl13b γ-Tubulin PH-Plcd1-GFP PH-Akt1-GFP

Reagents

Localization

References

α6F, monoclonal antibody, DSHB Antibody, Sigma Antibody, Santa Cruz, discontinued Custom antibody Phalloidin Monoclonal antibody clone 6-11B-1 (Sigma) Custom antibody Antibody, T5326 (Sigma-Aldrich) Transgene Transgene

Basolateral

Drummond et al. (1998)

Basal Basolateral

Gerlach and Wingert (2014) Duldulao et al. (2009)

Basolateral Apical Cilia

Duldulao et al. (2009)

Cilia Basal body

Sun et al. (2004) Tsujikawa and Malicki (2004)

PtdIns(4,5)P2 PtdIns(3,4,5)P3

Xu et al. (2017) Xu et al. (2017)

Tsujikawa and Malicki (2004)

3 Methods to analyze renal epithelial cells in the zebrafish pronephros

3.3.1.2 Protocol (1) Anesthetize embryos with 0.2% tricaine. Dilute formalin with PBST (1:2.7). Fix embryos in diluted formalin at room temperature (RT) for 1–2 h or at 4 °C overnight (ON). Alternatively fix with Dent’s fixative 20 °C ON. 50 zebrafish embryos can be easily handled in a 2 mL microcentrifuge tubes with 1 mL solution. For embryos older than 2 dpf, bleach pigment by incubating them with a solution of two parts methanol and one part 30% H2O2 at RT ON. (2) Replace with fresh methanol and the fixed embryos can now be stored at 20 °C. (3) Rehydrate embryos through 50% methanol in PBST for 5 min, followed by incubation in PBST for 5 min. (4) For embryos fixed with formalin, permeabilize with 1 mL of prechilled acetone, incubate at 20 °C for 7 min. Wash twice with PBST for 5 min each. (5) For embryos younger than 2 dfp, skip this step. Dilute 0.05% trypsin with PBST (1:20). Add diluted trypsin to embryos. Incubate at RT for 20–40 min. The optimal incubation time differs for embryos of different stages and batches of trypsin, therefore needs to be adjusted empirically. (6) Wash with 1 mL PBST 3 at RT for 5 min each. (7) Block in blocking buffer for half an hour. (8) Incubate with primary antibody in 200 μL blocking buffer for 2 h at RT or ON at 4 °C. (9) Wash with 1 mL PBST for 5 times, at least for 20 min each. (10) Incubate with conjugated secondary antibody in blocking buffer for 2 h at RT or ON at 4 °C. Shield sample from light with aluminum foil from this step on. For investigating the size and overall morphology of the tubule and duct or cilia bundle, chromogenic reaction is robust and does not require fluorescent microscope for visualization. Use HRP-conjugated secondary antibody and follow steps 11–14 for this method. Alternatively, fluorescent imaging provides better resolution essential for revealing subcellular structures. Use fluorophore-conjugated secondary antibodies and follow steps 11 and 15 for this method. (11) Wash with 1 mL PBST for 5  at RT, at least 20 min each. (12) Dissolve the 10 mg DAB tablet with 15 mL PBT and supplementing with 12 μL 30% hydrogen peroxide to make the working substrate solution. Incubate embryos in the working solution at RT. Monitor chromogenic reaction under a dissecting scope at increasing intervals. Avoid constant light exposure. In a successful experiment, a dark brown color will develop within 1 to 30 min. (13) Stop the reaction by washing in 1 mL PBST 3 5 min. Dehydrate completely with 1 mL methanol 2  5 min. Clear by incubating with BABB for 5 min and replace with fresh BABB. BABB is toxic. In addition, it dissolves tissue culture plates, although Eppendorf tubes (polypropylene) seems resistant. Always wear gloves, use glass pipette to handle samples and avoid contaminating working space and equipment. Dispose as hazardous waste according to local regulations.

177

178

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

(14) Transfer a few embryos onto to glass slide with a drop of BABB. Manually remove yolk with #5 watchmaker’s tweezers, flatten out embryos, remove excess BABB and yolk granules and cover with a cover slip. Observe using a compound microscope. Examples can be seen in Figs. 2C and 3C. In wild-type embryos the acetylated tubulin signal displays a linear pattern along the ventral side of the embryos, indicative of bundles of cilia from multi-ciliated cells within the duct lumen. In contrast, in the cilia biogenesis mutant arl13bhi459, a spotted pattern is seen (Fig. 2C). The mutant also shows enlarged diameter of the tubule labeled by α6F (Fig. 3C). (15) Transfer to a glass slide in PBST. Manually remove yolk, flatten out embryos. Remove PBST and yolk granules. Add mounting medium with DAPI. Cover with cover slip. Cure slides at RT ON protected from light. Observe using a fluorescent microscope.

3.3.2 Method 2. Cryosection and immunostaining to visualize structures of kidney epithelial cells Although technically more tedious and demanding, sections of embryos are more accessible for antibodies and provide better resolution for fine structure of cells. Here we provide a protocol for cryosection and immunostaining.

3.3.2.1 Materials • • • • •

Fixation and immunostaining reagents as in Method 1. Sucrose Tissue-Tek® OCT compound (Sakura) Dry ice CM1900 cryotome (Leica Microsystems, Buffalo Grove, IL).

3.3.2.2 Protocol (1) (2) (3) (4) (5) (6) (7) (8) (9) (10) (11)

Follow steps 1–3 in Method 1. Replace PBST with 10% (w/v) sucrose, incubate at RT for 1 h. Replace once with 20% (w/v) sucrose, incubate at RT for 1 h. Replace with 30% (w/v) sucrose and leave overnight at 4 °C. Transfer embryos to Tissue-Tek® OCT compound, orient embryos with a pipette tip or needle and freeze embryos in molds using dry ice. Cut in cross section at 8 μm using a cryotome. Collect sections on slides, dry slides at RT for 2 h. Slides can now be stored at 80 °C. Prior to immunolabeling, thaw slides at RT, rehydrated in PBST 3  5 min. Permeabilize with pre-chilled acetone for 7 min. Then wash 2  5 min in PBST. Block in blocking buffer for half an hour Incubate with primary antibody in 200 μL blocking buffer for 2 h at RT or ON at 4 °C. Wash with 1 mL PBST for 3 , for at least 10 min each.

References

(12) Incubate with your favorite secondary antibody in blocking buffer 2 h at RT or ON at 4 °C. Shield slides from light. (13) Wash with PBST for 3, at least for 10 min each. (14) Add mounting medium with DAPI, Cover with cover slip. Cure slides at RT ON protected from light. Observe under a fluorescent microscope.

4 Discussion and future direction In addition to gene inactivation, genetic variants could be pathogenic through other mechanisms such as acquiring dominant active or dominant negative functions. Orthologous models therefore could provide useful insight not revealed by null alleles. In contrast to indels, which now can be induced in targeted regions efficiently, precise replacement of specific genomic sequences is still incompatible with high throughput analysis. There are, however, exciting new technological improvements that may change this scenario (Gu, Posfai, & Rossant, 2018). Tagging of endogenous genes and tissue specific gene editing provide additional versatility for experimental designing. Although we focus on fixed samples in this chapter, the optical transparency makes zebrafish embryos a powerful system for live imaging. Multiple reporters, such as cilia markers, calcium indicators, calcium sponge and other biomarker, have been developed and can be used to address specific questions in vivo (Xu et al., 2017; Yuan et al., 2015). In short, zebrafish is a highly relevant genetic and in vivo model system for studying the kidney. Understanding its similarities and differences with the human kidney and available tools is essential to fully utilize its potential.

Acknowledgments Research in the Sun lab is supported by NIH (1R01DK113135, R01HL125885) and PKD foundation (213G16A).

References Aldahmesh, M. A., et al. (2014). IFT27, encoding a small GTPase component of IFT particles, is mutated in a consanguineous family with Bardet-Biedl syndrome. Human Molecular Genetics, 23(12), 3307–3315. Brinkman, E. K., et al. (2014). Easy quantitative assessment of genome editing by sequence trace decomposition. Nucleic Acids Research, 42(22), e168. Burger, A., et al. (2016). Maximizing mutagenesis with solubilized CRISPR-Cas9 ribonucleoprotein complexes. Development, 143(11), 2025–2037.

179

180

CHAPTER 10 In vivo analysis of renal epithelial cells in zebrafish

Carroll, T. J., & Das, A. (2011). Planar cell polarity in kidney development and disease. Organogenesis, 7(3), 180–190. Carroll, T. J., & Yu, J. (2012). The kidney and planar cell polarity. Current Topics in Developmental Biology, 101, 185–212. Chang-Panesso, M., & Humphreys, B. D. (2017). Cellular plasticity in kidney injury and repair. Nature Reviews. Nephrology, 13(1), 39–46. Combes, A. N., Davies, J. A., & Little, M. H. (2015). Cell-cell interactions driving kidney morphogenesis. Current Topics in Developmental Biology, 112, 467–508. Das, A., et al. (2013). Stromal-epithelial crosstalk regulates kidney progenitor cell differentiation. Nature Cell Biology, 15(9), 1035–1044. Drummond, I. (2002). The pronephros. Results and Problems in Cell Differentiation, 40, 322–345. Drummond, I. A., & Davidson, A. J. (2010). Zebrafish kidney development. Methods in Cell Biology, 100, 233–260. Drummond, I. A., et al. (1998). Early development of the zebrafish pronephros and analysis of mutations affecting pronephric function. Development, 125(23), 4655–4667. Duldulao, N. A., Lee, S., & Sun, Z. (2009). Cilia localization is essential for in vivo functions of the Joubert syndrome protein Arl13b/scorpion. Development, 136(23), 4033–4042. Gagnon, J. A., et al. (2014). Efficient mutagenesis by Cas9 protein-mediated oligonucleotide insertion and large-scale assessment of single-guide RNAs. PLoS One, 9(5), e98186. Gerlach, G. F., & Wingert, R. A. (2014). Zebrafish pronephros tubulogenesis and epithelial identity maintenance are reliant on the polarity proteins Prkc iota and zeta. Developmental Biology, 396(2), 183–200. Gu, B., Posfai, E., & Rossant, J. (2018). Efficient generation of targeted large insertions by microinjection into two-cell-stage mouse embryos. Nature Biotechnology, 36(7), 632–637. Hwang, W. Y., et al. (2013). Efficient genome editing in zebrafish using a CRISPR-Cas system. Nature Biotechnology, 31(3), 227–229. Jao, L. E., Wente, S. R., & Chen, W. (2013). Efficient multiplex biallelic zebrafish genome editing using a CRISPR nuclease system. Proceedings of the National Academy of Sciences of the United States of America, 110(34), 13904–13909. Jerman, S., & Sun, Z. (2017). Using zebrafish to study kidney development and disease. Current Topics in Developmental Biology, 124, 41–79. Karner, C. M., et al. (2009). Wnt9b signaling regulates planar cell polarity and kidney tubule morphogenesis. Nature Genetics, 41(7), 793–799. Kramann, R., et al. (2017). Gli1(+) pericyte loss induces capillary rarefaction and proximal tubular injury. Journal of the American Society of Nephrology: JASN, 28(3), 776–784. Kramer-Zucker, A. G., et al. (2005). Cilia-driven fluid flow in the zebrafish pronephros, brain and Kupffer’s vesicle is required for normal organogenesis. Development, 132(8), 1907–1921. Li, J., & Sun, Z. (2011). Qilin is essential for cilia assembly and normal kidney development in zebrafish. PLoS One, 6(11), e27365. Lienkamp, S. S., et al. (2012). Vertebrate kidney tubules elongate using a planar cell polaritydependent, rosette-based mechanism of convergent extension. Nature Genetics, 44(12), 1382–1387. Link, V., Shevchenko, A., & Heisenberg, C. P. (2006). Proteomics of early zebrafish embryos. BMC Developmental Biology, 6, 1.

References

Meng, X., et al. (2008). Targeted gene inactivation in zebrafish using engineered zinc-finger nucleases. Nature Biotechnology, 26(6), 695–701. Moreno-Mateos, M. A., et al. (2015). CRISPRscan: Designing highly efficient sgRNAs for CRISPR-Cas9 targeting in vivo. Nature Methods, 12(10), 982–988. Moreno-Mateos, M. A., et al. (2017). CRISPR-Cpf1 mediates efficient homology-directed repair and temperature-controlled genome editing. Nature Communications, 8(1), 2024. Olcese, C., et al. (2017). X-linked primary ciliary dyskinesia due to mutations in the cytoplasmic axonemal dynein assembly factor PIH1D3. Nature Communications, 8, 14279. Serluca, F. C., & Fishman, M. C. (2001). Pre-pattern in the pronephric kidney field of zebrafish. Development, 128(12), 2233–2241. Shah, A. N., et al. (2015). Rapid reverse genetic screening using CRISPR in zebrafish. Nature Methods, 12(6), 535–540. Soens, Z. T., et al. (2016). Hypomorphic mutations identified in the candidate Leber congenital amaurosis gene CLUAP1. Genetics in Medicine, 18(10), 1044–1051. Sun, Z., & Hopkins, N. (2001). vhnf1, the MODY5 and familial GCKD-associated gene, regulates regional specification of the zebrafish gut, pronephros, and hindbrain. Genes & Development, 15(23), 3217–3229. Sun, Z., et al. (2004). A genetic screen in zebrafish identifies cilia genes as a principal cause of cystic kidney. Development, 131(16), 4085–4093. Tsujikawa, M., & Malicki, J. (2004). Intraflagellar transport genes are essential for differentiation and survival of vertebrate sensory neurons. Neuron, 42(5), 703–716. Vasilyev, A., et al. (2009). Collective cell migration drives morphogenesis of the kidney nephron. PLoS Biology, 7(1), e9. Verdeguer, F., et al. (2010). A mitotic transcriptional switch in polycystic kidney disease. Nature Medicine, 16(1), 106–110. Westerfield, M. (1993). The zebrafish book: A guide for the laboratory use of zebrafish (Brachydanio rerio). Eugene, OR: The University of Oregon Press. Wingert, R. A., & Davidson, A. J. (2008). The zebrafish pronephros: A model to study nephron segmentation. Kidney International, 73(10), 1120–1127. Xu, W., et al. (2017). The Joubert syndrome protein Inpp5e controls ciliogenesis by regulating phosphoinositides at the apical membrane. Journal of the American Society of Nephrology: JASN, 28(1), 118–129. Yuan, S., & Sun, Z. (2009). Microinjection of mRNA and morpholino antisense oligonucleotides in zebrafish embryos. Journal of Visualized Experiments(27). Yuan, S., et al. (2015). Intraciliary calcium oscillations initiate vertebrate left-right asymmetry. Current Biology, 25(5), 556–567.

181