Chapter 9 Functional Analysis of Hox Genes in Zebrafish Franck Ladam and Charles G. Sagerström Abstract The zebrafish model organism is well suited to study the role of specific genes, such as hox genes, in embryogenesis and organ function. The ability to modulate the activity of hox genes in living zebrafish embryos represents a cornerstone of such functional analyses. In this chapter we outline the basic methodology for nucleic acid injections into 1–2-cell-stage zebrafish embryos. We also report variations in this method to allow injection of mRNA, DNA, and antisense oligonucleotides to either overexpress, knock down, or knock out specific genes in zebrafish embryos. Key words hox, Zebrafish, Microinjection, Over-expression, Morpholino, Tol2, Transgenesis, Zinc-finger nuclease, TALE nuclease, CRISPR-Cas system
1
Introduction hox genes act as regulators of transcription in development and disease and have been studied for decades using various model organisms or cell lines. The zebrafish (Danio rerio) expresses 48 Hox proteins and has been widely used to decipher their function during embryonic development. For instance, hox genes control anteroposterior and dorsoventral patterning of the embryo and are involved in more subtle events such as neuronal migration and specification (reviewed in [1]). The zebrafish is a useful model to study key developmental mechanisms as its embryos are easy to obtain, easy to grow, and transparent. Moreover, its genome is fully sequenced and 69 % of the genes have at least one human ortholog [2]. In this methods chapter we describe some of the techniques available to study hox gene function in zebrafish. The main method relies on the injection of nucleic acid into 1–2-cellstage embryos (adapted from the zebrafish handbook [3]) to either over-express, knock down, or knock out specific hox genes during embryonic development. We describe (1) a general protocol for injection into 1–2-cell-stage embryos and (2) specific protocols to efficiently produce and inject nucleic acids in order to manipulate
Yacine Graba and René Rezsohazy (eds.), Hox Genes: Methods and Protocols, Methods in Molecular Biology, vol. 1196, DOI 10.1007/978-1-4939-1242-1_9, © Springer Science+Business Media New York 2014
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and study hox gene function in zebrafish. Injected embryos can be subsequently used for classical molecular biology, cellular biology, or biochemistry experiments (e.g., Western blot, in situ hybridization, immunostaining, chromatin immunoprecipitation, RT-PCR). Finally, even though the focus of this chapter is on hox gene function, all the described techniques are suitable for the study of any given gene.
2
Materials
2.1 General Laboratory Materials and Reagents
1. RNAse/DNAse-free DEPC-treated water.
2.2 Fish Crosses, Embryo Injections, and Growth
1. Agarose.
2. 1.7 ml low-binding microcentrifuge tubes (Corning). 3. 50, 65, and 95 °C dry block heaters.
2. 10 cm diameter plastic petri dishes. 3. Injection mold with six ramps, one 90° and one 45° beveled side (Adaptive Science Tools). 4. 0.5 % Red phenol solution (Sigma-Aldrich). 5. Borosilicate glass capillaries 1.0 OD × 0.5 mm ID/Fiber (FHC). 6. pc-10 needle puller (Narishige). 7. 1.0 l crossing tanks with dividers (Aquaneering). 8. Microloader pipet tips, for prefilling microinjection capillaries. 9. A pair of sharp stainless dissecting forceps. 10. MZ6 modular stereomicroscope with 6.3:1 zoom and eyepiece graticule (Leica) (Fig. 1). 11. Micro-manipulator M3301 (World Precision Instrument) (Fig. 1).
Fig. 1 Photograph of microinjection setup. Equipment required for nucleic acid injections into zebrafish embryos is labeled
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12. Magnetic stand (World Precision Instrument) (Fig. 1). 13. PLI90 pneumatic pico-injector (Harvard Apparatus) (Fig. 1). 14. Pipet pump. 15. Mineral oil. 16. Plain microscope glass slides. 17. 146 mm Borosilicate glass Pasteur pipettes (Thermo Fisher Scientific). 18. Methylene blue water to grow zebrafish embryos: 0.0002 % methylene blue, 0.006 % instant ocean salts (Instant Ocean), milli-Q water. 19. Fine mesh strainer diameter 5 cm (Fante’s). 20. 28.5 °C Lab-Line Imperial III incubator (Thermo Fisher Scientific). 2.3 mRNA Synthesis, MO Preparation
1. mMessage mMachine in vitro transcription kit (Ambion). 2. QIAquick PCR purification kit (Qiagen). 3. NanoDrop Scientific).
3300
fluorospectrometer
(Thermo
Fisher
4. RNA loading dye (Ambion). 5. 1× Tris–Borate–EDTA buffer: 220 mM Tris, 180 mM boric acid, 4 mM EDTA. 2.4 Tol2 Transgenesis
1. pTol2 kit containing Gateway-compatible vectors and the transposase-encoding vector PCS2FA-transposase [4, 5]. The kit is freely available from the Chien Lab [5]. 2. Sterile Luria Broth bacteria medium: 25 g of powder in 1 l of milli-Q water. 3. NotI restriction enzyme. 4. HiSpeed Plasmid Midi Kit (Qiagen). 5. 25× Tricaine solution: 15.3 mM tricaine (Sigma-Aldrich), 20 mM Tris–HCl pH 9.0. 6. Dissection spring scissor (Roboz). 7. 0.2 ml PCR tubes. 8. Lysis buffer: 10 mM Tris–HCl pH 7.5, 0.5 % sodium dodecyl sulfate, 100 μg/ml Proteinase K. 9. Phenol–chloroform–isoamyl alcohol solution. 10. 3 M Sodium acetate pH 5.2. 11. 70 and 100 % ethanol. 12. Chloroform solution. 13. One Taq Hot Start DNA polymerase (New England Biolabs). 14. 6× DNA loading dye (New England Biolabs).
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Methods
3.1 General Protocol for Microinjection in Zebrafish
The following section describes general guidelines to achieve efficient injection into 1–2-cell-stage zebrafish embryos. Steps 1–6 are performed on the day before injection. 1. Prepare the injection plates by pouring 15 ml of 1 % agarose melted in methylene blue water into a 10 cm plastic dish and set the injection mold on top of the hot agarose. Let the agarose solidify and remove the mold. 2. Adjust settings on the pc-10 needle puller (see Note 1). 3. Tightly set a borosilicate capillary on the needle puller. The center of the capillary should face the electric resistance filament to obtain two needles of equal size. 4. Press the start button. As the electric resistance filament warms up, one end of the capillary is pulled down to obtain two injection needles. 5. Prepare the solution to inject (see Subheadings 3.2–3.4) by mixing the desired amount of material (for instance 1 μg of mRNA), 1 μl of 0.5 % red phenol solution, and RNAse/ DNAse-free water to a 10 μl final volume into a 1.7 ml microcentrifuge tube (see Note 2). 6. At the end of the day, place zebrafish at a ratio of two males for three females into the inner chamber of a crossing tank filled with fish water, using the divider to keep males and females separated. Leave the fish overnight at 28 °C in the dark (see Note 3). 7. The following morning as the light comes on, mix the males and females together in the inner chamber filled with fresh water. As eggs are laid and fertilized they will sink to the bottom of the outer chamber of the tank (see Note 4). 8. Using a p2 pipet and Microloader pipet tips, backload 2 μl of the injection solution (from step 5) into the injection needle. 9. Set the filled needle on the needle holder. Cut off the very tip of the needle using a thin pair of dissecting forceps, monitoring the process under the stereomicroscope (see Note 5). 10. Press the footswitch of the pico-injector and make sure that a drop of red liquid is forming at the tip of the needle. 11. Quantify the amount of material contained in a drop by injecting into 100 μl of mineral oil set on a microscope glass slide. Measure the radius of the drop with the calibrated graticule on the stereomicroscope. Use the formula 4/3πr3 to determine the volume of the sphere and therefore the volume injected. Using the volume of the drop and the original concentration of your solution, calculate the amount of material contained in one drop. 12. Adjust the pressure and duration parameters on the pico-injector to obtain the drop size you need for your experiment (Fig. 1).
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Fig. 2 Schematic illustrating injection into one-cell-stage zebrafish embryos. (a) Fertilized one-cell-stage embryos are placed on the agarose injection-mold ramps. (b) Close-up of zebrafish embryos showing the injection needle and the injected solution (red dot). Note that the needle needs to get through the chorion and the cell membrane before injecting the solution. (c) Side view of a one-cell-stage zebrafish embryo set on an agarose mold ramp
13. Collect the embryos into a 10 cm plastic dish using a mesh strainer and a squirt bottle filled with methylene blue egg water (Fig. 2, see Note 6). 14. Using a pipette, place the collected embryos on the injection plate and orient them on the ramps using forceps or a p200 pipet tip so that the cell is to the top (Fig. 2, see Note 7). 15. Carefully introduce the needle into the cell and inject the desired amount of material by pressing the footswitch. Make sure that the red stain remains in the cell when pulling the needle out of the embryo (Fig. 2). 16. Move the needle to the next embryo and repeat step 15 to inject as many embryos as needed (see Note 8). 17. Place the injected embryos into a 10 cm plastic dish containing methylene blue water and incubate at 28 °C until reaching the desired developmental stage (see Note 9). 3.2
mRNA Injections
Exogenous gene expression in zebrafish can be accomplished by injecting 5′ 7-methyl guanosine capped messenger RNAs (mRNAs) directly into 1–2-cell-stage embryos. Capped mRNA in vitro transcription necessitates the prior cloning of the coding cDNA of interest into an expression vector allowing for in vitro transcription by RNA polymerases SP6, T3, or T7. Capped mRNAs mimic endogenous mRNAs and, unlike expression plasmids, allow the expression of the transgene throughout the embryo (see Notes 10 and 11). Injection of capped mRNA can also be used to express
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non-endogenous proteins such as those needed for genome engineering methods (e.g., zinc-finger nucleases and TALE nucleases, see Subheading 3.5). 1. Linearize 10 μg of the plasmid containing the cDNA with a restriction enzyme cutting at the 3′ end of the coding sequence. 2. Run 500 ng of the linearized plasmid on a gel composed of 1 % agarose melted in 1× Tris–borate–EDTA buffer and check for complete digestion. 3. Purify the linearized plasmid using the PCR purification kit and elute in 30 μl of RNase/DNase-free milli-Q water. 4. Quantify by loading 2 μl of the purified plasmid on the nanodrop spectrophotometer. 5. Run an in vitro transcription reaction on 2 μg of the digested DNA, following the mMessage mMachine in vitro transcription kit instructions. 6. Digest the DNA template by adding 2 μl of RNase-free DNase to the reaction (provided in the mMessage kit). 7. Purify the capped mRNAs using the RNAeasy RNA purification kit and elute in 30 μl of RNase/DNase-free water. 8. Quantify by loading 2 μl of the purified capped mRNA onto a nanodrop spectrophotometer. 9. Mix 300 ng of the capped mRNAs with water and RNA loading dye to a 1× final concentration, heat up to 65 °C for 5 min, and set on ice for another 5 min. 10. Load on a gel composed of 1 % agarose melted in 1× Tris– borate–EDTA buffer to assess the quality of the RNA sample (see Note 12). 3.3 Morpholino Antisense Oligonucleotide Injections
Morpholino antisense oligonucleotides (MOs) consist of nucleic acid bases bound to morpholine rings instead of deoxyribose rings [6]. They function by steric blockade of mRNA-dependent processes such as splicing and translation. MOs are usually synthesized to be complementary to mRNA translation start site or to exon–intron boundaries (see Note 13). MOs are very stable and easy to work with and sequences can be designed and synthesized at will by the Gene Tools company (Gene Tools, LLC). They are usually provided as a white dried powder. 1. Resuspend the MO powder in sterile milli-Q water to make a 3 mM stock solution (see Note 14). 2. Make a 1 mM MO solution ready for injections by mixing 3 μl of 3 mM MO, 1 μl 0.5 % red phenol dye, and 5 μl of milli-Q water in a 1.7 ml tube (see Note 15). 3. Heat up the MO solution to 65 °C for 5 min and set on ice for another 5 min.
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3.4 DNA Injections for Transgenesis
As discussed earlier, mRNA injection into zebrafish embryos allows widespread, but transient, expression of a specific gene. However, detailed analyses of gene function may require driving expression in a time- and tissue-restricted fashion. This can be achieved through the expression of a transgene under the control of a tissuespecific promoter/enhancer region. Yet, plasmid injection generates low, transient, and mosaic expression of the transgene and is therefore often not suitable for gene studies. Thus, the following section describes a simple protocol to achieve efficient and stable transgene integration into the zebrafish genome using the Tol2 system. The Tol2 system allows the transposition into the genome of a transgene flanked by sequences (called 5′ and 3′ arms) derived from the Tol2 transposable element (originally identified in medaka fish) when co-injected with an mRNA expressing a transposase into 1–2-cell-stage zebrafish embryos [7–10]. This method efficiently integrates large DNA fragments and, in the case of hox genes, has been successfully used to insert a 70 kb DNA fragment containing the pufferfish (Fugu rubripes) hoxAa cluster (hoxa5 through hoxa13) into the zebrafish genome [11].
3.4.1 Preparation of the pTol2-Transgene Vector
The plasmid to be injected should consist of the transgene cloned into the pTol2 vector (referred below as pTol2-transgene plasmid). 1. Purify the pTol2-transgene plasmid, from exponentially growing bacterial culture in Luria Broth medium, using the QiaQuick Midi prep Kit. Elute the plasmid in milli-Q water. Do not use the EB solution provided in the kit (see Note 16). 2. Quantify by loading 2 μl of the purified plasmid onto a nanodrop spectrophotometer.
3.4.2 Preparation of the Transposase mRNA
1. Digest the PCS2FA-transposase vector with the NotI restriction enzyme. 2. Purify the digested plasmid and in vitro transcribe the transposase mRNA as described in Subheading 3.2.
3.4.3 Injection into 1–2-Cell-Stage Embryos
1. Mix 1 μg of pTol2-transgene with 1 μg of the transposase mRNA, 1 μl of 0.5 % red phenol solution, and RNAse/DNAsefree water to a 10 μl final volume into a 1.7 ml microcentrifuge tube (see Note 17). 2. Inject into 1–2-cell-stage embryos.
3.4.4 Validation of Transgene Insertion
Validation of the transgene insertion into the genome can be achieved by polymerase chain reaction (PCR) amplification of purified zebrafish genomic DNA (see Note 18). This method is performed on injected embryos grown to the adult stage [12]. 1. In a small plastic box, place two or three fish into tricaine diluted in fish water to a 1× concentration. Wait until the fish stop swimming, but gill movements persist.
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2. Collect one fish with a plastic spoon and place it on a 10 cm plastic dish. 3. Hold the very tip of the caudal fin with a pair of forceps, clip a small piece of fin using a pair of dissection spring scissors, place in a 0.2 ml PCR tube, and proceed to the next fish (see Note 19). 4. Add 100 μl of lysis buffer and incubate for 4 h at 50 °C (see Note 20). 5. Vortex briefly. 6. Incubate the samples at 95 °C for 10 min to inactivate the Proteinase K contained in the lysis buffer. 7. Pipette 50 μl of the lysate into a new 1.7 ml tube and add 150 μl of milli-Q water. 8. Add 400 μl of phenol–chloroform–isoamyl alcohol solution and vortex vigorously. 9. Spin at 16,000 × g for 15 min at 4 °C. 10. Pipette the aqueous top layer into a new 1.7 ml tube, add 1 volume of chloroform, and vortex vigorously. 11. Spin at 16,000 × g for 5 min at room temperature. 12. Pipette the aqueous top layer into a new 1.7 ml tube. 13. Add 0.1 volume of 3 M pH 5.2 sodium acetate and 2 volumes of 100 % ethanol. 14. Place the samples at −20 °C for 30 min. 15. Centrifuge at 16,000 × g for 15 min at 4 °C. 16. Wash the DNA pellet with 1 ml of 70 % ethanol. 17. Centrifuge at 16,000 × g for 5 min at 4 °C. 18. Dry the pellet by leaving the tube open at room temperature for 5–10 min. 19. Resuspend the DNA pellet in 50 μl milli-Q water. 20. Run a PCR reaction by mixing the following in a 0.2 ml PCR tube (see Note 21): –
5 μl of 5× One Taq standard reaction buffer.
–
0.5 μl of 10 mM dNTPs.
–
0.5 μl of 10 μM forward primer.
–
0.5 μl of 10 μM reverse primer.
–
0.125 μl of One Taq Hot Start DNA polymerase.
–
100 ng of purified genomic DNA.
–
Nuclease-free water to 25 μl.
21. Mix 15 μl of the PCR product with 3 μl of 6× DNA loading dye and load on a gel composed of 1 or 2 % agarose melted in 1× Tris–borate–EDTA buffer.
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3.5 Genome Engineering Technologies
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As mentioned previously, the use of MOs for loss-of-function experiments has various drawbacks. Within the past few years, three different methods—zinc-finger nucleases (ZFNs), TALE nucleases (TALENs), and the CRISPR-Cas system—have been employed to allow specific and stable disruption of zebrafish genes. These technologies involve ectopic expression, in the embryo, of a nuclease moiety targeted to the gene of interest by a sequencespecific recognition module (detailed descriptions of each technique are available elsewhere [13–15]). Gene disruption by ZFNs and TALENs is achieved by injecting mRNA (as in Subheading 3.2) encoding a chimeric protein composed of the FokI nuclease fused to the recognition module (a zinc finger or a TALE protein, respectively). Mutagenesis by the CRISPR-Cas system requires the injection of a guide RNA and mRNA encoding the Cas9 nuclease (as in Subheading 3.2). Identification of mutant founders is then accomplished using PCR and/or sequencing analogously to how transgenic founders are identified (Subheading 3.4.4). These techniques have been used in zebrafish to knock out genes by generating mutations (insertions or deletions) but also to introduce specific DNA sequences (knock-in) or induce chromosomal deletions or inversion [16–18].
Notes 1. Recommended settings are provided by the manufacturer [19]. In our experiments we routinely use the following settings: STEP1, N°1 heater set to 69.5 °C, tensile force corresponding to 4 weights. 2. The solution can be used right away or frozen at −80 °C for later use. Avoid freeze-thaw cycles. 3. Using the crossing tanks described here, a ratio of two males for three females gives good yields of embryos; however other ratios might be considered to increase the number of embryos. 4. It is important to keep the adults separated from the embryos, as adults will eventually feed on the embryos. 5. The size and shape of the needle tip is a critical parameter. It should be as thin as possible but still be able to get through the chorion. 6. Collect the embryos before the first mitotic division occurs (i.e., before 35–45 min post-fertilization). 7. You can also place the embryos so that the cell is to one side (Fig. 2). We do not recommend working with the cell under the yolk as this may result in clogging of the needle. 8. As you inject embryos keep a steady drop size by adjusting the pico-injector parameters.
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9. Regularly remove dead embryos to avoid growth of bacteria and mold. 10. Plasmid DNA injection in zebrafish embryos results in mosaic expression of the transgene. Moreover, since protein synthesis requires transcription and translation inside the cells, the expression of the transgene takes longer than when mRNA is injected. For those reasons RNA injections are more suitable for transient overexpression of a specific gene in zebrafish embryos. 11. Injection of mRNA might induce nonspecific effects due to nonphysiological expression levels of the protein. Therefore, it is recommended to determine the optimal amount of mRNA to inject by doing a titration experiment and choosing the lowest concentration of mRNA that produces a specific phenotype. Moreover, we recommend injecting a nonfunctional proteincoding RNA as a control condition. Also, because the injection is made at the one-cell stage, the mRNA of interest may be expressed at places and times the endogenous mRNA is not. In the case of Hox proteins, this is of particular importance as their endogenous pattern and timing of expression are tightly regulated during development (e.g., the first hox gene (hoxb1b) is expressed 6 h post-fertilization and is restricted to the hindbrain and the most posterior part of the embryo [20–22]). Hence, misexpression of Hox proteins may lead to homeotic transformations [21–24], often resulting in defective embryonic development. Therefore other methods to study hox function should also be considered (see Subheadings 3.3 and 3.4). 12. While this does not allow for determining the actual size of the mRNA, it is a simple way to assess for RNA integrity and detect any potential DNA or RNA contaminants. 13. The use of MOs to study gene function in zebrafish has several drawbacks. (a) MOs may have nonspecific effects and several controls should be employed to confirm the biological significance of any MO-based phenotype [25]. First, one should consider the use of a nonfunctional MO. Such MOs usually have mutations introduced into the sequence of the original MO so that its target recognition is altered. Second, off-target effects may result in activation of a p53-dependent apoptotic response. Therefore, co-injection of the gene-specific MO with a p53 targeting MO should be used to determine if an apoptotic phenotype is p53 mediated. Third, a titration experiment should be done in order to determine the optimal amount of MO leading to a specific phenotype and minimal death of the embryos. Fourth, a rescue experiment should be performed by co-injecting the MO and the target mRNA (modified to be unaffected by the MO). (b) The efficiency of the knockdown is often hard to assess due to the lack of available antibodies to study protein expression in zebrafish. (c) The effect of MOs is transient. (d) MOs are comparatively expensive.
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14. Keep at −20 °C. 15. MO concentration and volume can be modified according to your needs. 16. Quality of the plasmid is a critical parameter, as poor-quality DNA preparation leads to embryo death. 17. The relative amounts of pTol2-transgene vector and PCS2FAtransposase mRNA should be experimentally determined to obtain an efficient integration of the transgene into the genome. 18. Several copies of the transgene are often found on different chromosomes throughout the genome and the number of transgene copies therefore decreases as the transgenic fish are outbred to a wild-type background. Moreover, transgene insertion is random and may result in gene disruption. Thus, to avoid nonspecific effects and ensure steady transgene expression, one should consider a few recommendations: (a) outbreed the transgenic fish to a wild-type background through several generations; (b) determine the transgene copy number by southern blot [9]; (c) localize the transgene insertions in the genome [9]; and (d) raise and work with at least two different transgenic lines. 19. Do not cut too close to the body as you would hurt the fish. A 3 mm-by-3 mm piece of fin is usually enough to obtain sufficient amounts of DNA. 20. Samples can also be incubated overnight at 50 °C. 21. Optimal PCR parameters (i.e., annealing temperature, duration of elongation, number of cycles) depend upon the primers and the sequences amplified. Primers should allow specific amplification of the transgene. If the gene to be overexpressed is found endogenously in the genome one might consider using primers located within sequences specific to the Tol2 construct.
Acknowledgements This work was supported by NIH grants NS038183 and HD065081 to CGS. References 1. Alexander T, Nolte C, Krumlauf R (2009) Hox genes and segmentation of the hindbrain and axial skeleton. Annu Rev Cell Dev Biol 25:431–456 2. Howe K, Clark MD, Torroja CF et al (2013) The zebrafish reference genome sequence and its relationship to the human genome. Nature 496:498–503
3. Westerfield M (2007) The zebrafish book, 5th edition; a guide for the laboratory use of zebrafish (Danio rerio). University of Oregon Press, Eugene 4. Villefranc JA, Amigo J, Lawson ND (2007) Gateway compatible vectors for analysis of gene function in the zebrafish. Dev Dyn 236: 3077–3087
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5. Kwan KM, Fujimoto E, Grabher C et al (2007) The Tol2kit: a multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev Dyn 236:3088–3099 6. Ekker SC, Larson JD (2001) Morphant technology in model developmental systems. Genesis 30:89–93 7. Kawakami K (2007) Tol2: a versatile gene transfer vector in vertebrates. Genome Biol 8(Suppl 1):S7 8. Suster ML, Abe G, Schouw A et al (2011) Transposon-mediated BAC transgenesis in zebrafish. Nat Protoc 6:1998–2021 9. Suster ML, Kikuta H, Urasaki A et al (2009) Transgenesis in zebrafish with the Tol2 transposon system. In: Cartwright EJ (ed) Methods in molecular biology, vol 561. Humana Press, Totowa, NJ, pp 41–63 10. Kikuta H, Kawakami K (2009) Transient and stable transgenesis using Tol2 transposon vectors. In: Lieschke GJ, Oates AC, Kawakami K. (eds) Methods in molecular biology, vol 546. Humana Press, Totowa, NJ, pp 69–84 11. Suster ML, Sumiyama K, Kawakami K (2009) Transposon-mediated BAC transgenesis in zebrafish and mice. BMC Genomics 10:477 12. Kimmel CB, Ballard WW, Kimmel SR et al (1995) Stages of embryonic development of the zebrafish. Dev Dyn 203:253–310 13. Urnov FD, Rebar EJ, Holmes MC et al (2010) Genome editing with engineered zinc finger nucleases. Nat Rev Genet 11:636–646 14. Gaj T, Gersbach CA, Barbas CF III (2013) ZFN, TALEN, and CRISPR/Cas-based methods for genome engineering. Trends Biotechnol 31:397–405 15. Hwang WY, Fu Y, Reyon D, Maeder ML et al (2013) Efficient genome editing in zebrafish using a CRISPR-Cas system. Nat Biotechnol 31:227–229
16. Xiao A, Wang Z, Hu Y et al (2013) Chromosomal deletions and inversions mediated by TALENs and CRISPR/Cas in zebrafish. Nucleic Acids Res. doi:10.1093/ nar/gkt464 17. Gupta A, Hall VL, Kok FO et al (2013) Targeted chromosomal deletions and inversions in zebrafish. Genome Res 23:1008–1017 18. Bedell VM, Wang Y, Campbell JM et al (2012) In vivo genome editing using a high-efficiency TALEN system. Nature 491:114–118 19. Narishige Web News (2007) http://news.narishige-group.com/pdf/news001en.pdf . Accessed 4 Dec 2013 20. Alexandre D, Clarke JD, Oxtoby E et al (1996) Ectopic expression of Hoxa-1 in the zebrafish alters the fate of the mandibular arch neural crest and phenocopies a retinoic acid-induced phenotype. Development 122:735–746 21. Vlachakis N, Choe SK, Sagerström CG (2001) Meis3 synergizes with Pbx4 and Hoxb1b in promoting hindbrain fates in the zebrafish. Development 128:1299–1312 22. McClintock JM, Carlson R, Mann DM et al (2001) Consequences of Hox gene duplication in the vertebrates: an investigation of the zebrafish Hox paralogue group 1 genes. Development 128:2471–2484 23. Choe S-K, Zhang X, Hirsch N et al (2011) A screen for hoxb1-regulated genes identifies ppp1r14al as a regulator of the rhombomere 4 Fgf-signaling center. Dev Biol. doi:10.1016/j. ydbio.2011.05.676 24. Bruce AE, Oates AC, Prince VE et al (2001) Additional hox clusters in the zebrafish: divergent expression patterns belie equivalent activities of duplicate hoxB5 genes. Evol Dev 3:127–144 25. Gerety SS, Wilkinson DG (2011) Morpholino artifacts provide pitfalls and reveal a novel role for pro-apoptotic genes in hindbrain boundary development. Dev Biol 350:279–289
1
Introduction hox genes act as regulators of transcription in development and disease and have been studied for decades using various model organisms or cell lines. The zebrafish (Danio rerio) expresses 48 Hox proteins and has been widely used to decipher their function during embryonic development. For instance, hox genes control anteroposterior and dorsoventral patterning of the embryo and are involved in more subtle events such as neuronal migration and specification (reviewed in [1]). The zebrafish is a useful model to study key developmental mechanisms as its embryos are easy to obtain, easy to grow, and transparent. Moreover, its genome is fully sequenced and 69 % of the genes have at least one human ortholog [2]. In this methods chapter we describe some of the techniques available to study hox gene function in zebrafish. The main method relies on the injection of nucleic acid into 1–2-cellstage embryos (adapted from the zebrafish handbook [3]) to either over-express, knock down, or knock out specific hox genes during embryonic development. We describe (1) a general protocol for injection into 1–2-cell-stage embryos and (2) specific protocols to efficiently produce and inject nucleic acids in order to manipulate
Yacine Graba and René Rezsohazy (eds.), Hox Genes: Methods and Protocols, Methods in Molecular Biology, vol. 1196, DOI 10.1007/978-1-4939-1242-1_9, © Springer Science+Business Media New York 2014
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and study hox gene function in zebrafish. Injected embryos can be subsequently used for classical molecular biology, cellular biology, or biochemistry experiments (e.g., Western blot, in situ hybridization, immunostaining, chromatin immunoprecipitation, RT-PCR). Finally, even though the focus of this chapter is on hox gene function, all the described techniques are suitable for the study of any given gene.
2
Materials
2.1 General Laboratory Materials and Reagents
1. RNAse/DNAse-free DEPC-treated water.
2.2 Fish Crosses, Embryo Injections, and Growth
1. Agarose.
2. 1.7 ml low-binding microcentrifuge tubes (Corning). 3. 50, 65, and 95 °C dry block heaters.
2. 10 cm diameter plastic petri dishes. 3. Injection mold with six ramps, one 90° and one 45° beveled side (Adaptive Science Tools). 4. 0.5 % Red phenol solution (Sigma-Aldrich). 5. Borosilicate glass capillaries 1.0 OD × 0.5 mm ID/Fiber (FHC). 6. pc-10 needle puller (Narishige). 7. 1.0 l crossing tanks with dividers (Aquaneering). 8. Microloader pipet tips, for prefilling microinjection capillaries. 9. A pair of sharp stainless dissecting forceps. 10. MZ6 modular stereomicroscope with 6.3:1 zoom and eyepiece graticule (Leica) (Fig. 1). 11. Micro-manipulator M3301 (World Precision Instrument) (Fig. 1).
Fig. 1 Photograph of microinjection setup. Equipment required for nucleic acid injections into zebrafish embryos is labeled
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12. Magnetic stand (World Precision Instrument) (Fig. 1). 13. PLI90 pneumatic pico-injector (Harvard Apparatus) (Fig. 1). 14. Pipet pump. 15. Mineral oil. 16. Plain microscope glass slides. 17. 146 mm Borosilicate glass Pasteur pipettes (Thermo Fisher Scientific). 18. Methylene blue water to grow zebrafish embryos: 0.0002 % methylene blue, 0.006 % instant ocean salts (Instant Ocean), milli-Q water. 19. Fine mesh strainer diameter 5 cm (Fante’s). 20. 28.5 °C Lab-Line Imperial III incubator (Thermo Fisher Scientific). 2.3 mRNA Synthesis, MO Preparation
1. mMessage mMachine in vitro transcription kit (Ambion). 2. QIAquick PCR purification kit (Qiagen). 3. NanoDrop Scientific).
3300
fluorospectrometer
(Thermo
Fisher
4. RNA loading dye (Ambion). 5. 1× Tris–Borate–EDTA buffer: 220 mM Tris, 180 mM boric acid, 4 mM EDTA. 2.4 Tol2 Transgenesis
1. pTol2 kit containing Gateway-compatible vectors and the transposase-encoding vector PCS2FA-transposase [4, 5]. The kit is freely available from the Chien Lab [5]. 2. Sterile Luria Broth bacteria medium: 25 g of powder in 1 l of milli-Q water. 3. NotI restriction enzyme. 4. HiSpeed Plasmid Midi Kit (Qiagen). 5. 25× Tricaine solution: 15.3 mM tricaine (Sigma-Aldrich), 20 mM Tris–HCl pH 9.0. 6. Dissection spring scissor (Roboz). 7. 0.2 ml PCR tubes. 8. Lysis buffer: 10 mM Tris–HCl pH 7.5, 0.5 % sodium dodecyl sulfate, 100 μg/ml Proteinase K. 9. Phenol–chloroform–isoamyl alcohol solution. 10. 3 M Sodium acetate pH 5.2. 11. 70 and 100 % ethanol. 12. Chloroform solution. 13. One Taq Hot Start DNA polymerase (New England Biolabs). 14. 6× DNA loading dye (New England Biolabs).
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Methods
3.1 General Protocol for Microinjection in Zebrafish
The following section describes general guidelines to achieve efficient injection into 1–2-cell-stage zebrafish embryos. Steps 1–6 are performed on the day before injection. 1. Prepare the injection plates by pouring 15 ml of 1 % agarose melted in methylene blue water into a 10 cm plastic dish and set the injection mold on top of the hot agarose. Let the agarose solidify and remove the mold. 2. Adjust settings on the pc-10 needle puller (see Note 1). 3. Tightly set a borosilicate capillary on the needle puller. The center of the capillary should face the electric resistance filament to obtain two needles of equal size. 4. Press the start button. As the electric resistance filament warms up, one end of the capillary is pulled down to obtain two injection needles. 5. Prepare the solution to inject (see Subheadings 3.2–3.4) by mixing the desired amount of material (for instance 1 μg of mRNA), 1 μl of 0.5 % red phenol solution, and RNAse/ DNAse-free water to a 10 μl final volume into a 1.7 ml microcentrifuge tube (see Note 2). 6. At the end of the day, place zebrafish at a ratio of two males for three females into the inner chamber of a crossing tank filled with fish water, using the divider to keep males and females separated. Leave the fish overnight at 28 °C in the dark (see Note 3). 7. The following morning as the light comes on, mix the males and females together in the inner chamber filled with fresh water. As eggs are laid and fertilized they will sink to the bottom of the outer chamber of the tank (see Note 4). 8. Using a p2 pipet and Microloader pipet tips, backload 2 μl of the injection solution (from step 5) into the injection needle. 9. Set the filled needle on the needle holder. Cut off the very tip of the needle using a thin pair of dissecting forceps, monitoring the process under the stereomicroscope (see Note 5). 10. Press the footswitch of the pico-injector and make sure that a drop of red liquid is forming at the tip of the needle. 11. Quantify the amount of material contained in a drop by injecting into 100 μl of mineral oil set on a microscope glass slide. Measure the radius of the drop with the calibrated graticule on the stereomicroscope. Use the formula 4/3πr3 to determine the volume of the sphere and therefore the volume injected. Using the volume of the drop and the original concentration of your solution, calculate the amount of material contained in one drop. 12. Adjust the pressure and duration parameters on the pico-injector to obtain the drop size you need for your experiment (Fig. 1).
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Fig. 2 Schematic illustrating injection into one-cell-stage zebrafish embryos. (a) Fertilized one-cell-stage embryos are placed on the agarose injection-mold ramps. (b) Close-up of zebrafish embryos showing the injection needle and the injected solution (red dot). Note that the needle needs to get through the chorion and the cell membrane before injecting the solution. (c) Side view of a one-cell-stage zebrafish embryo set on an agarose mold ramp
13. Collect the embryos into a 10 cm plastic dish using a mesh strainer and a squirt bottle filled with methylene blue egg water (Fig. 2, see Note 6). 14. Using a pipette, place the collected embryos on the injection plate and orient them on the ramps using forceps or a p200 pipet tip so that the cell is to the top (Fig. 2, see Note 7). 15. Carefully introduce the needle into the cell and inject the desired amount of material by pressing the footswitch. Make sure that the red stain remains in the cell when pulling the needle out of the embryo (Fig. 2). 16. Move the needle to the next embryo and repeat step 15 to inject as many embryos as needed (see Note 8). 17. Place the injected embryos into a 10 cm plastic dish containing methylene blue water and incubate at 28 °C until reaching the desired developmental stage (see Note 9). 3.2
mRNA Injections
Exogenous gene expression in zebrafish can be accomplished by injecting 5′ 7-methyl guanosine capped messenger RNAs (mRNAs) directly into 1–2-cell-stage embryos. Capped mRNA in vitro transcription necessitates the prior cloning of the coding cDNA of interest into an expression vector allowing for in vitro transcription by RNA polymerases SP6, T3, or T7. Capped mRNAs mimic endogenous mRNAs and, unlike expression plasmids, allow the expression of the transgene throughout the embryo (see Notes 10 and 11). Injection of capped mRNA can also be used to express
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non-endogenous proteins such as those needed for genome engineering methods (e.g., zinc-finger nucleases and TALE nucleases, see Subheading 3.5). 1. Linearize 10 μg of the plasmid containing the cDNA with a restriction enzyme cutting at the 3′ end of the coding sequence. 2. Run 500 ng of the linearized plasmid on a gel composed of 1 % agarose melted in 1× Tris–borate–EDTA buffer and check for complete digestion. 3. Purify the linearized plasmid using the PCR purification kit and elute in 30 μl of RNase/DNase-free milli-Q water. 4. Quantify by loading 2 μl of the purified plasmid on the nanodrop spectrophotometer. 5. Run an in vitro transcription reaction on 2 μg of the digested DNA, following the mMessage mMachine in vitro transcription kit instructions. 6. Digest the DNA template by adding 2 μl of RNase-free DNase to the reaction (provided in the mMessage kit). 7. Purify the capped mRNAs using the RNAeasy RNA purification kit and elute in 30 μl of RNase/DNase-free water. 8. Quantify by loading 2 μl of the purified capped mRNA onto a nanodrop spectrophotometer. 9. Mix 300 ng of the capped mRNAs with water and RNA loading dye to a 1× final concentration, heat up to 65 °C for 5 min, and set on ice for another 5 min. 10. Load on a gel composed of 1 % agarose melted in 1× Tris– borate–EDTA buffer to assess the quality of the RNA sample (see Note 12). 3.3 Morpholino Antisense Oligonucleotide Injections
Morpholino antisense oligonucleotides (MOs) consist of nucleic acid bases bound to morpholine rings instead of deoxyribose rings [6]. They function by steric blockade of mRNA-dependent processes such as splicing and translation. MOs are usually synthesized to be complementary to mRNA translation start site or to exon–intron boundaries (see Note 13). MOs are very stable and easy to work with and sequences can be designed and synthesized at will by the Gene Tools company (Gene Tools, LLC). They are usually provided as a white dried powder. 1. Resuspend the MO powder in sterile milli-Q water to make a 3 mM stock solution (see Note 14). 2. Make a 1 mM MO solution ready for injections by mixing 3 μl of 3 mM MO, 1 μl 0.5 % red phenol dye, and 5 μl of milli-Q water in a 1.7 ml tube (see Note 15). 3. Heat up the MO solution to 65 °C for 5 min and set on ice for another 5 min.
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3.4 DNA Injections for Transgenesis
As discussed earlier, mRNA injection into zebrafish embryos allows widespread, but transient, expression of a specific gene. However, detailed analyses of gene function may require driving expression in a time- and tissue-restricted fashion. This can be achieved through the expression of a transgene under the control of a tissuespecific promoter/enhancer region. Yet, plasmid injection generates low, transient, and mosaic expression of the transgene and is therefore often not suitable for gene studies. Thus, the following section describes a simple protocol to achieve efficient and stable transgene integration into the zebrafish genome using the Tol2 system. The Tol2 system allows the transposition into the genome of a transgene flanked by sequences (called 5′ and 3′ arms) derived from the Tol2 transposable element (originally identified in medaka fish) when co-injected with an mRNA expressing a transposase into 1–2-cell-stage zebrafish embryos [7–10]. This method efficiently integrates large DNA fragments and, in the case of hox genes, has been successfully used to insert a 70 kb DNA fragment containing the pufferfish (Fugu rubripes) hoxAa cluster (hoxa5 through hoxa13) into the zebrafish genome [11].
3.4.1 Preparation of the pTol2-Transgene Vector
The plasmid to be injected should consist of the transgene cloned into the pTol2 vector (referred below as pTol2-transgene plasmid). 1. Purify the pTol2-transgene plasmid, from exponentially growing bacterial culture in Luria Broth medium, using the QiaQuick Midi prep Kit. Elute the plasmid in milli-Q water. Do not use the EB solution provided in the kit (see Note 16). 2. Quantify by loading 2 μl of the purified plasmid onto a nanodrop spectrophotometer.
3.4.2 Preparation of the Transposase mRNA
1. Digest the PCS2FA-transposase vector with the NotI restriction enzyme. 2. Purify the digested plasmid and in vitro transcribe the transposase mRNA as described in Subheading 3.2.
3.4.3 Injection into 1–2-Cell-Stage Embryos
1. Mix 1 μg of pTol2-transgene with 1 μg of the transposase mRNA, 1 μl of 0.5 % red phenol solution, and RNAse/DNAsefree water to a 10 μl final volume into a 1.7 ml microcentrifuge tube (see Note 17). 2. Inject into 1–2-cell-stage embryos.
3.4.4 Validation of Transgene Insertion
Validation of the transgene insertion into the genome can be achieved by polymerase chain reaction (PCR) amplification of purified zebrafish genomic DNA (see Note 18). This method is performed on injected embryos grown to the adult stage [12]. 1. In a small plastic box, place two or three fish into tricaine diluted in fish water to a 1× concentration. Wait until the fish stop swimming, but gill movements persist.
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2. Collect one fish with a plastic spoon and place it on a 10 cm plastic dish. 3. Hold the very tip of the caudal fin with a pair of forceps, clip a small piece of fin using a pair of dissection spring scissors, place in a 0.2 ml PCR tube, and proceed to the next fish (see Note 19). 4. Add 100 μl of lysis buffer and incubate for 4 h at 50 °C (see Note 20). 5. Vortex briefly. 6. Incubate the samples at 95 °C for 10 min to inactivate the Proteinase K contained in the lysis buffer. 7. Pipette 50 μl of the lysate into a new 1.7 ml tube and add 150 μl of milli-Q water. 8. Add 400 μl of phenol–chloroform–isoamyl alcohol solution and vortex vigorously. 9. Spin at 16,000 × g for 15 min at 4 °C. 10. Pipette the aqueous top layer into a new 1.7 ml tube, add 1 volume of chloroform, and vortex vigorously. 11. Spin at 16,000 × g for 5 min at room temperature. 12. Pipette the aqueous top layer into a new 1.7 ml tube. 13. Add 0.1 volume of 3 M pH 5.2 sodium acetate and 2 volumes of 100 % ethanol. 14. Place the samples at −20 °C for 30 min. 15. Centrifuge at 16,000 × g for 15 min at 4 °C. 16. Wash the DNA pellet with 1 ml of 70 % ethanol. 17. Centrifuge at 16,000 × g for 5 min at 4 °C. 18. Dry the pellet by leaving the tube open at room temperature for 5–10 min. 19. Resuspend the DNA pellet in 50 μl milli-Q water. 20. Run a PCR reaction by mixing the following in a 0.2 ml PCR tube (see Note 21): –
5 μl of 5× One Taq standard reaction buffer.
–
0.5 μl of 10 mM dNTPs.
–
0.5 μl of 10 μM forward primer.
–
0.5 μl of 10 μM reverse primer.
–
0.125 μl of One Taq Hot Start DNA polymerase.
–
100 ng of purified genomic DNA.
–
Nuclease-free water to 25 μl.
21. Mix 15 μl of the PCR product with 3 μl of 6× DNA loading dye and load on a gel composed of 1 or 2 % agarose melted in 1× Tris–borate–EDTA buffer.
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3.5 Genome Engineering Technologies
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As mentioned previously, the use of MOs for loss-of-function experiments has various drawbacks. Within the past few years, three different methods—zinc-finger nucleases (ZFNs), TALE nucleases (TALENs), and the CRISPR-Cas system—have been employed to allow specific and stable disruption of zebrafish genes. These technologies involve ectopic expression, in the embryo, of a nuclease moiety targeted to the gene of interest by a sequencespecific recognition module (detailed descriptions of each technique are available elsewhere [13–15]). Gene disruption by ZFNs and TALENs is achieved by injecting mRNA (as in Subheading 3.2) encoding a chimeric protein composed of the FokI nuclease fused to the recognition module (a zinc finger or a TALE protein, respectively). Mutagenesis by the CRISPR-Cas system requires the injection of a guide RNA and mRNA encoding the Cas9 nuclease (as in Subheading 3.2). Identification of mutant founders is then accomplished using PCR and/or sequencing analogously to how transgenic founders are identified (Subheading 3.4.4). These techniques have been used in zebrafish to knock out genes by generating mutations (insertions or deletions) but also to introduce specific DNA sequences (knock-in) or induce chromosomal deletions or inversion [16–18].
Notes 1. Recommended settings are provided by the manufacturer [19]. In our experiments we routinely use the following settings: STEP1, N°1 heater set to 69.5 °C, tensile force corresponding to 4 weights. 2. The solution can be used right away or frozen at −80 °C for later use. Avoid freeze-thaw cycles. 3. Using the crossing tanks described here, a ratio of two males for three females gives good yields of embryos; however other ratios might be considered to increase the number of embryos. 4. It is important to keep the adults separated from the embryos, as adults will eventually feed on the embryos. 5. The size and shape of the needle tip is a critical parameter. It should be as thin as possible but still be able to get through the chorion. 6. Collect the embryos before the first mitotic division occurs (i.e., before 35–45 min post-fertilization). 7. You can also place the embryos so that the cell is to one side (Fig. 2). We do not recommend working with the cell under the yolk as this may result in clogging of the needle. 8. As you inject embryos keep a steady drop size by adjusting the pico-injector parameters.
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9. Regularly remove dead embryos to avoid growth of bacteria and mold. 10. Plasmid DNA injection in zebrafish embryos results in mosaic expression of the transgene. Moreover, since protein synthesis requires transcription and translation inside the cells, the expression of the transgene takes longer than when mRNA is injected. For those reasons RNA injections are more suitable for transient overexpression of a specific gene in zebrafish embryos. 11. Injection of mRNA might induce nonspecific effects due to nonphysiological expression levels of the protein. Therefore, it is recommended to determine the optimal amount of mRNA to inject by doing a titration experiment and choosing the lowest concentration of mRNA that produces a specific phenotype. Moreover, we recommend injecting a nonfunctional proteincoding RNA as a control condition. Also, because the injection is made at the one-cell stage, the mRNA of interest may be expressed at places and times the endogenous mRNA is not. In the case of Hox proteins, this is of particular importance as their endogenous pattern and timing of expression are tightly regulated during development (e.g., the first hox gene (hoxb1b) is expressed 6 h post-fertilization and is restricted to the hindbrain and the most posterior part of the embryo [20–22]). Hence, misexpression of Hox proteins may lead to homeotic transformations [21–24], often resulting in defective embryonic development. Therefore other methods to study hox function should also be considered (see Subheadings 3.3 and 3.4). 12. While this does not allow for determining the actual size of the mRNA, it is a simple way to assess for RNA integrity and detect any potential DNA or RNA contaminants. 13. The use of MOs to study gene function in zebrafish has several drawbacks. (a) MOs may have nonspecific effects and several controls should be employed to confirm the biological significance of any MO-based phenotype [25]. First, one should consider the use of a nonfunctional MO. Such MOs usually have mutations introduced into the sequence of the original MO so that its target recognition is altered. Second, off-target effects may result in activation of a p53-dependent apoptotic response. Therefore, co-injection of the gene-specific MO with a p53 targeting MO should be used to determine if an apoptotic phenotype is p53 mediated. Third, a titration experiment should be done in order to determine the optimal amount of MO leading to a specific phenotype and minimal death of the embryos. Fourth, a rescue experiment should be performed by co-injecting the MO and the target mRNA (modified to be unaffected by the MO). (b) The efficiency of the knockdown is often hard to assess due to the lack of available antibodies to study protein expression in zebrafish. (c) The effect of MOs is transient. (d) MOs are comparatively expensive.
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14. Keep at −20 °C. 15. MO concentration and volume can be modified according to your needs. 16. Quality of the plasmid is a critical parameter, as poor-quality DNA preparation leads to embryo death. 17. The relative amounts of pTol2-transgene vector and PCS2FAtransposase mRNA should be experimentally determined to obtain an efficient integration of the transgene into the genome. 18. Several copies of the transgene are often found on different chromosomes throughout the genome and the number of transgene copies therefore decreases as the transgenic fish are outbred to a wild-type background. Moreover, transgene insertion is random and may result in gene disruption. Thus, to avoid nonspecific effects and ensure steady transgene expression, one should consider a few recommendations: (a) outbreed the transgenic fish to a wild-type background through several generations; (b) determine the transgene copy number by southern blot [9]; (c) localize the transgene insertions in the genome [9]; and (d) raise and work with at least two different transgenic lines. 19. Do not cut too close to the body as you would hurt the fish. A 3 mm-by-3 mm piece of fin is usually enough to obtain sufficient amounts of DNA. 20. Samples can also be incubated overnight at 50 °C. 21. Optimal PCR parameters (i.e., annealing temperature, duration of elongation, number of cycles) depend upon the primers and the sequences amplified. Primers should allow specific amplification of the transgene. If the gene to be overexpressed is found endogenously in the genome one might consider using primers located within sequences specific to the Tol2 construct.
Acknowledgements This work was supported by NIH grants NS038183 and HD065081 to CGS. References 1. Alexander T, Nolte C, Krumlauf R (2009) Hox genes and segmentation of the hindbrain and axial skeleton. Annu Rev Cell Dev Biol 25:431–456 2. Howe K, Clark MD, Torroja CF et al (2013) The zebrafish reference genome sequence and its relationship to the human genome. Nature 496:498–503
3. Westerfield M (2007) The zebrafish book, 5th edition; a guide for the laboratory use of zebrafish (Danio rerio). University of Oregon Press, Eugene 4. Villefranc JA, Amigo J, Lawson ND (2007) Gateway compatible vectors for analysis of gene function in the zebrafish. Dev Dyn 236: 3077–3087
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5. Kwan KM, Fujimoto E, Grabher C et al (2007) The Tol2kit: a multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev Dyn 236:3088–3099 6. Ekker SC, Larson JD (2001) Morphant technology in model developmental systems. Genesis 30:89–93 7. Kawakami K (2007) Tol2: a versatile gene transfer vector in vertebrates. Genome Biol 8(Suppl 1):S7 8. Suster ML, Abe G, Schouw A et al (2011) Transposon-mediated BAC transgenesis in zebrafish. Nat Protoc 6:1998–2021 9. Suster ML, Kikuta H, Urasaki A et al (2009) Transgenesis in zebrafish with the Tol2 transposon system. In: Cartwright EJ (ed) Methods in molecular biology, vol 561. Humana Press, Totowa, NJ, pp 41–63 10. Kikuta H, Kawakami K (2009) Transient and stable transgenesis using Tol2 transposon vectors. In: Lieschke GJ, Oates AC, Kawakami K. (eds) Methods in molecular biology, vol 546. Humana Press, Totowa, NJ, pp 69–84 11. Suster ML, Sumiyama K, Kawakami K (2009) Transposon-mediated BAC transgenesis in zebrafish and mice. BMC Genomics 10:477 12. Kimmel CB, Ballard WW, Kimmel SR et al (1995) Stages of embryonic development of the zebrafish. Dev Dyn 203:253–310 13. Urnov FD, Rebar EJ, Holmes MC et al (2010) Genome editing with engineered zinc finger nucleases. Nat Rev Genet 11:636–646 14. Gaj T, Gersbach CA, Barbas CF III (2013) ZFN, TALEN, and CRISPR/Cas-based methods for genome engineering. Trends Biotechnol 31:397–405 15. Hwang WY, Fu Y, Reyon D, Maeder ML et al (2013) Efficient genome editing in zebrafish using a CRISPR-Cas system. Nat Biotechnol 31:227–229
16. Xiao A, Wang Z, Hu Y et al (2013) Chromosomal deletions and inversions mediated by TALENs and CRISPR/Cas in zebrafish. Nucleic Acids Res. doi:10.1093/ nar/gkt464 17. Gupta A, Hall VL, Kok FO et al (2013) Targeted chromosomal deletions and inversions in zebrafish. Genome Res 23:1008–1017 18. Bedell VM, Wang Y, Campbell JM et al (2012) In vivo genome editing using a high-efficiency TALEN system. Nature 491:114–118 19. Narishige Web News (2007) http://news.narishige-group.com/pdf/news001en.pdf . Accessed 4 Dec 2013 20. Alexandre D, Clarke JD, Oxtoby E et al (1996) Ectopic expression of Hoxa-1 in the zebrafish alters the fate of the mandibular arch neural crest and phenocopies a retinoic acid-induced phenotype. Development 122:735–746 21. Vlachakis N, Choe SK, Sagerström CG (2001) Meis3 synergizes with Pbx4 and Hoxb1b in promoting hindbrain fates in the zebrafish. Development 128:1299–1312 22. McClintock JM, Carlson R, Mann DM et al (2001) Consequences of Hox gene duplication in the vertebrates: an investigation of the zebrafish Hox paralogue group 1 genes. Development 128:2471–2484 23. Choe S-K, Zhang X, Hirsch N et al (2011) A screen for hoxb1-regulated genes identifies ppp1r14al as a regulator of the rhombomere 4 Fgf-signaling center. Dev Biol. doi:10.1016/j. ydbio.2011.05.676 24. Bruce AE, Oates AC, Prince VE et al (2001) Additional hox clusters in the zebrafish: divergent expression patterns belie equivalent activities of duplicate hoxB5 genes. Evol Dev 3:127–144 25. Gerety SS, Wilkinson DG (2011) Morpholino artifacts provide pitfalls and reveal a novel role for pro-apoptotic genes in hindbrain boundary development. Dev Biol 350:279–289
