Combining genotypic and phenotypic analyses on single mutant zebrafish larvae

Graphical abstract


G R A P H I C A L A B S T R A C T A B S T R A C T
Zebrafish is a powerful animal model used to study vertebrate embryogenesis, organ development and diseases (Gut et al., 2017) [1]. The usefulness of the model was established as a result of various large forward genetic screens identifying mutants in almost every organ or cell type (Driever et al., 1996;Haffter et al., 1996) [2,3]. More recently, the advent of genome editing methodologies, including TALENs (Sander et al., 2011) [4] and the CRISPR/ Cas9 technology (Hwang et al., 2013) [5], led to an increase in the production of zebrafish mutants. A number of these mutations are homozygous lethal at the embryonic or larval stages preventing the generation of homozygous mutant zebrafish lines. Here, we present a method allowing both genotyping and phenotype analyses of mutant zebrafish larvae from heterozygous zebrafish incrosses. The procedure is based on the genotyping of the larval tail after transection, whereas phenotypic studies are performed on the anterior part of the zebrafish larvae.

Subject area
Biochemistry, Genetics and Molecular Biology More specific subject area Zebrafish developmental biology Method name Single zebrafish larvae characterization Name and reference of original method Resource availability

Method details
Forwardand reverse genetics in zebrafish has established a huge variety of mutants [1][2][3][4][5]. Since numerous mutations are homozygous lethal at early developmental stages, methods allowing both genotyping and phenotype analyses of mutant zebrafish embryos and larvae are required.The method is designed to investigate the phenotype of sibling zebrafish larvae from heterozygous zebrafish incrosses. As so, single zebrafish larvae must be genotyped in order to eventually establish a correlation between different phenotypes and distinct genotypes. After transection, the larval tail is used for genotyping, whereas phenotypic studies are performed on the anterior part of the zebrafish larvae. 1. Anesthetize zebrafish embryos or larvae with the MS-222 anesthetic solution in 6-well plates. 2. Sedated embryos or larvae are transferred using a P20 micropipettor equipped with a cut-off tip into a 10-cm Petri dish containing MS-222 anesthetic solution 3. Under a stereomicroscope, cut the tail with a sterile scalpel distal to the end of the intestine in order to preserve most of the embryo for phenotypic analyses and to recover a tail biopsy allowing easy handling and efficient DNA extraction (Fig. 1A). 4. Transfer the tail biopsy into a 0.2 mL sterile tube for DNA extraction whereas the anterior part of the larvae is transferred into a 24-well dish containing the MS-222 overdose solution for euthanasia and placed on ice

Note
After 5-10 min in a MS-222 anesthetic solution bath, zebrafish embryos and larvae are fully sedated. Embryos and larvae may be kept in the anesthetic solution up to 2 h without effect on viability.
DNA extraction from tail biopsies and genotyping DNA extraction from zebrafish embryonic and larval tails is achieved using a protocol based on the use of sodium hydroxide and Tris, modified from Meeker et al. [6]. This method allows fast genomic DNA extraction used for genotyping by RFLP assay (Restriction Fragment Length Polymorphism; [7]). The method detects a mutation that either creates or abolishes a site recognized by a specific restriction enzyme. In the RFLP assay, a sequence of interest is first PCRamplified and the PCR product is subjected to restriction enzyme digestion to identify whether the amplified alleles are wild-type or mutant.    Under these conditions, it is possible to distinguish wild-type and ezh2 ul2 alleles by RFLP [8].

Notes
This procedure may also be used on paraformaldehyde-fixed samples to genotype entire embryos and larvae in 20 mL 50 mM NaOH after in situ hybridization, Alcian blue-Alizarin red staining or Oil red-O staining [8,9]. The setting of the PCR conditions may be modified according to the annealing temperature of the Forward and Reverse primers.
In step 5, a master mix (N + 1) of all reagents could be made and 17.5 mL of the mix subsequently added to the DNA. In step 7, a master mix (N + 1) containing all restriction reagents may be used to ensure that all samples are equally proceeded. Since the identification of mutant alleles relies on the absence of a restriction site, including a wildtype control in the genotyping analysis may be a useful positive control. The protocol details a genotyping strategy based on RFLP, but alternate methods such as derived Cleaved Amplification Polymorphic Sequence (dCAPS; [10]) or High Resolution Melting Analysis (HRMA; [11]) genotyping assays could be used after the DNA extraction procedure.

Histological analysis
For histological studies, sections of the anterior part of zebrafish embryos or larvae are performed after paraffin embedding.

Equipment
Glass pillboxes (Dutscher, # 211672) Hybridization oven Heating plate Stainless steel molds (Dutscher, # 040731) Paraffin fountain (Leica, # 1120) Microtome (Leica, # RM2245, or equivalent) Cover slips (WWR, # 631-1575) Glass slides, SuperFrost plus (Thermo, # 4951 PLUS4) Embryos and larvae are fixed overnight at 4 C 3. Wash the embryos or larvae once with PBS for 5 min 4. Transfer the embryos and larvae into glass pillboxes 5. Dehydrate the embryos and larvae with successive dilutions of ethanol in water: 10 min in 1.5 mL 30% (vol/vol) ethanol, 10 min in 1.5 mL 50% (vol/vol) ethanol, 10 min in 1.5 mL 70% (vol/vol) ethanol, 15 min in 1.5 mL 100% ethanol 6. Put the embryos and larvae for 10 min in 1.5 mL 50% ethanol -50% Claral (vol/vol) at room temperature 7. Incubate embryos and larvae for 15 min in 1.5 mL 100% Claral at room temperature 8. After removal of the Claral, fill the glass pillboxes with pre-warmed paraffin in a paraffin fountain overnight at 58 C, and incubate 15 min in an oven at 58 C 9. Change the paraffin and incubate again for 15 min at 58 C 10. Put the embryos and larvae in new warm paraffin solution into stainless steel molds and orientate the samples as required 11. Place the molds on the heating plate at 58 C for 30 min. Then, on the heating plate at 37 C for 1 h, and finally at room temperature for 4 h to overnight 12. Place the molds for at least 4 h at 4 C before demolding and keep the zebrafish-containing paraffin blocks at 4 C 13. Reduce the paraffin around the embryo with a razor blade 14. Stick the zebrafish-containing paraffin block to a bigger one (Fig. 1C) which is placed on the microtome 15. Perform 5 mm sections with the microtome 16. 16. Deposit the paraffin sections in a water drop onto a glass slide placed on a heating plate at 37 C for 5 min and then remove the water 17. Place the glass slide on the heating plate at 37 C for 30 min to 1 h maximum 18. Store the glass slides for at least 1 night at 37 C in an incubator Histological sections could then be analyzed by hematoxylin-eosin staining (Fig. 1C), TUNEL labeling, in situ hybridization, or in situ immunohistochemistry [8].

Protein analysis by western blot
For protein studies (Fig. 1D), the anterior part of zebrafish embryos or larvae is transferred in a 1.5 mL Eppendorf tube, snap frozen in liquid nitrogen and kept at À80 C for at least 1 night. Since epigenetic regulations, including histone modifications, play a crucial role in vertebrate development, the method describes both total protein and histone extraction procedures. This method allows the recovery of approximately 3 mg histone proteins per 7 dpf embryo.
If several embryos or larvae from the same genotype are processed together, mash 5-10 embryos in 15 mL TEB, resuspend in 7.5 mL TEB after the first centrifugation (step 5) and extract the histones with 7.5 mL 0.2 N HCl (step 7). Protein concentration is measured using the Bradford reagent (Biorad, # 500 00 06) with 2 mL of sample.
Western blot procedure 1. Boil the samples 10 min at 95 C 2. Load the protein samples onto a SDS denaturating gel (NuPAGE, 4-12% Bis-Tris polyacrylamide gel, Invitrogen) and run for about 90 min at constant 150 V 3. Transfer the proteins to a nitrocellulose membrane by iBlot Gel Transfer Stacks (Invitrogen) at 20 V for 7 min (preset program P3) 4. Block the membrane by incubation in Blocking Buffer, 1 h on an orbital shaker. Alternatively, membranes could be blocked overnight at 4 C 5. Incubate the membrane with the primary antibody in Blocking Buffer for 1 h at room temperature. The incubation of the membrane with the antibody is performed by spreading the antibody solution with a pipette on the membrane, placed onto a Parafilm M flattened on the bench 6. Wash the membrane 3 times 10 min in PBST 7. Incubate the membrane with the secondary antibody in Blocking Buffer for 1 h at room temperature 8. Wash the membrane 3 times 10 min in PBST 9. Visualize the protein using a chemiluminescence substrate (Super Signal West Femto, Thermo) and a Luminescent Image Analyzer (LAS-4000, Fujifilm).

Note
The Western blot results strongly rely on the quality of the antibodies. The limited number of zebrafish antibodies available often makes protein-based analyses more difficult. Fig. 1D shows Western blots revealing the actin protein from total protein extracts and histone H3 from histone extraction from the anterior part of a single wild-type zebrafish larvae at 9 dpf. The primary antibodies used were rabbit anti-actin (1:10,000; A2066, Sigma) and rabbit anti-H3 (1:5000; ab1791, Abcam). The secondary antibody was a peroxidase conjugated donkey anti-rabbit antibody (1:10,000; 711-035-152, Jackson ImmunoResearch).
Membrane stripping for re-use 1. Incubate the membrane 2 times 10 min at room temperature in freshly prepared Stripping Buffer for total proteins 2. Wash the membrane twice 5 min in PBST at room temperature 3. Block the membrane 1 h in Blocking Buffer at room temperature, before re-use with a Primary Antibody For histone blots the procedure is modified as follow: 1. Incubate the membrane 30 min in an hybridization oven at 50 C with pre-warmed Stripping Buffer for histone blots 2. Shake the membrane every 10 min 3. Wash the membrane 10 times 5 min in PBST at room temperature 4. Block the membrane 1 h in Blocking Buffer at room temperature before re-use with a Primary Antibody

RNA analysis
For RNA studies (Fig. 1E), the anterior part of zebrafish embryos or larvae is transferred in a 1.5 mL Eppendorf tube, 200 mL Trizol is added before snap freezing in liquid nitrogen and storage at À80 C for at least 1 night. The RNAs are extracted based on a modified version of the protocol from de Jong et al. [12].

Note
The Reverse transcription procedure follows the instruction provided with the SuperScriptIII Kit (Invitrogen).
PCR procedure 1. Transfer 5 mL of cDNAs from the reverse transcription into a 0.2 mL PCR tube 2. Add 2.5 mL of 10Â Polymerase Buffer, 0.5 mL 10 mM dNTPs Mix, 2.5 mL 25 mM MgCl 2 , 0.5 mL of each forward and reverse primers (10 mM), 0.5 mL Taq DNA Polymerase (5 U/mL) and 13 mL distilled water (final volume is 25 mL)

Notes
The PCR conditions may be modified according to the annealing temperature of the Forward and Reverse primers.
In step 2, a master mix (N + 1) of all reagents could be made and 20 mL of the mix subsequently added to the cDNA samples. Fig. 1E shows the expression of actin revealed by RT-PCR from RNAs extracted from the anterior part of a single wild-type zebrafish larvae at 9 dpf. PCR amplification of the actin cDNA was performed using forward (5 0 -CGTGACATCAAGGAGAAGCT-3 0 ) and reverse (5 0 -ATCCACATCTGCTGGAAGGT-3 0 ) generates a 442 bp DNA fragment.

Study of caudal spinal cord regeneration
The ability to regenerate tissues after amputation is part of the phenotypic studies that can be performed on zebrafish mutants at the larval stage [8].

Materials
Living zebrafish embryos at 3 dpf  2D shows the genotyping of the progeny of an ezh2 ul2 [8] heterozygous incross using tail biopsy from 3 dpf embryos generated by this procedure.