We thank Dr. Jennifer Emtage for critical reading of the manuscript, Ryan Gill for technical assistance and help with zebrafish care and Dr. Stephen C. Ekker (Mayo Clinic, Rochster, Minnesota) for reagents. Our work has been supported by Temple University College of Science and Technology and the National Institutes of Health (R01-HD061749).

1. Production of the F0 Generation by Microinjection of the Gene Trap.
We have found that embryos of different genetic backgrounds display different tolerances to this procedure, with pet-store-based wild type and Tuebingen - Long Fin (TLF) being the most tolerant while AB and Tuebingen strains have poorer survival.
<ol>
	<li>Preparation of transposon DNA. Prepare GBT-B1 (pDB783, Balciuniene et al. in preparation) plasmid DNA using standard QIAprep miniprep kit (Qiagen 27106). It is essential to include the PB wash step (which is optional in QIAprep procedure), otherwise the miniprep DNA will be contaminated by RNase A, leading to degradation of the transposase mRNA. Before injection, dilute the DNA in RNase-free water (Ambion AM9937) to 10 ng/&mu;l.</li>
	<li>Preparation of transposase mRNA. Linearize pT3TS/Tol2 (pDB600) 13 using XbaI and transcribe the linearized DNA using a mMessage Machine T3 (Ambion AM1348) in vitro transcription kit. We have modified the in vitro transcription reaction by adding 0.5 &mu;l of RiboLock RNase inhibitor (ThermoFisher Fermentas EO0381). After the DNase treatment step, purify mRNA using an RNeasy MinElute kit (Qiagen 74204). Assess the quality of the in vitro transcribed mRNA by agarose gel electrophoresis. Dilute the mRNA in RNase-free water to 40 ng/&mu;l, and store as 2 &mu;l aliquots at -80 &deg;C.</li>
	<li>Preparation of the microinjection mix. Before injection, add 8 &mu;l of diluted transposon DNA to a 2 &mu;l aliquot of transposase mRNA.</li>
	<li>Microinjection. Inject 3 nl of DNA/RNA mixture into the yolk of 1 cell zebrafish embryos using standard microinjection techniques, aiming for the yolk/blastomere interface. Collect embryos every 20 min to ensure injection at 1 cell stage. In our experience, a skillful person can easily inject 1,000 embryos in 1-1.5 hr.</li>
	<li>After injection, run 5 &mu;l of leftover injection solution on a 1% agarose gel for quality control, to ensure that the transposase mRNA has not been degraded.</li>
	<li>Injected embryo care. In late afternoon, remove unfertilized and abnormal embryos and distribute embryos to 60-80 per 100 mm Petri dish. Abnormal and dead embryos are removed at 1 day post fertilization (dpf), 2 dpf and 3 dpf.</li>
	<li>Screening for GFP expression. Screen embryos without overt defects for GFP fluorescence at 3 dpf (Figure 3). Screening can be done either on a stereomicroscope equipped with fluorescence (we use Nikon a SMZ1500) or on an upright microscope (we use a Zeiss AxioImager with a 5X Fluar objective). Low GFP expression indicates either unsuccessful injection or degradation of transposase RNA due to RNase contamination (Figure 3A). Select about 30% of the brightest embryos which either have GFP expression in multiple tissues or in a high percentage of cells of a specific tissue (compare Figures 3B and 3C). From an injection of 1,000 embryos, we usually have up to 100 developmentally-normal, high-GFP-expressing embryos.</li>
	<li>Raise selected F0 embryos using standard zebrafish rearing procedures. It is reasonable to expect that 20-30% of the injected embryos selected for rearing will survive to adulthood.</li>
</ol>
2. Maintenance of the UAS:mRFP Tester Line.
We have generated a 14xUAS:mRFP line marked by lens-specific &gamma;Cry:GFP. Since UAS is subject to silencing through DNA methylation, it is important to select fish with low levels of silencing to propagate the stock.
<ol>
	<li>Set up individual UAS:mRFP homozygotes for mating with one of our most reliable Gal4 gene trap lines, nsftpl6 (Balciuniene et al. in preparation).</li>
	<li>On the next day, transfer the UAS:mRFP fish which gave embryos to individual static tanks, while the embryos are collected and cleaned.</li>
	<li>Screen embryos for GFP and RFP fluorescence at 1 dpf and 3 dpf.</li>
	<li>Intercross fish which gave embryos with high mRFP expression to establish the next generation of UAS:mRFP line.</li>
</ol>
3. Screening of the F0 Fish.
<ol>
	<li>Cross individual F0 with the homozygous UAS:mRFP fish (Figure 4). We usually inject our GBTs into lines with wild-type or leopard pigmentation patterns, and our homozygous UAS:mRFP line is in brass background. Therefore GBT-injected F0 fish can be readily distinguished from the UAS:mRFP fish, eliminating the need to record whether the F0 being screened was a male or a female as well as potential errors resulting from mistakes in recording and/or determining the sex of each fish. Personnel with very limited experience, such us undergraduate students, are therefore able to set up and break down fish matings. This approach has a disadvantage that two different genetic backgrounds are being used, potentially complicating interpretation of loss of function phenotypes.</li>
	<li>On the next day, return the brass UAS:mRFP fish to their tank. Place the F0 fish which gave embryos into individual static tanks (we use Hagen Exo Terra Faunarium Small, but any small tank can be used for this purpose) while embryos are collected and screened.</li>
	<li>Clean and transfer collected embryos into new Petri dishes (&lt;100 per dish) in the afternoon of day 0.</li>
	<li>Screen embryos for GFP and RFP fluorescence at 1 dpf, 2 dpf and 3 dpf. Pull embryos positive for both GFP and RFP out, sort them by GFP/RFP expression pattern (if more than one pattern is observed in a clutch) and raise them to establish F1 generation.</li>
	<li>Euthanize the F0 fish which produced only negative embryos (out of a total number of 50-100 embryos).</li>
	<li>Cross the F0 fish which produced GFP/RFP-positive progeny again to get a second batch of positives.</li>
</ol>
4. Screening of the F1 Fish.
The F1 fish are screened to establish F2 families, to document gene trap expression pattern and to freeze batches of embryos for molecular identification of gene trap loci by 5' RACE and inverse PCR.
<ol>
	<li>Cross individual F1 fish with the homozygous UAS:mRFP fish (Figure 4).</li>
	<li>Next day, place the F1 fish that gave embryos into individual static tanks while the embryos are collected and screened.</li>
	<li>Screen embryos for GFP and RFP fluorescence at 1 dpf, 2 dpf and 3 dpf. Separate and photograph embryos positive for both GFP and RFP 24.</li>
	<li>At 5 dpf, freeze batches of 20 GFP/RFP-positive and 20 GFP/RFP-negative embryos for identification of insertionally mutated genes by inverse PCR and 5' RACE 9,10. Raise remaining GFP/RFP-positive embryos to establish F2 generation.</li>
</ol>

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These authors have nothing to disclose.

The gene trap vectors and methodology described here are already being used in several independent laboratories. In our experience, there are two critical steps in the process. The first critical step is to achieve high rates of gene trap integration in injected F0 embryos. Failure at this step can often be attributed to RNase contamination of the injection reagents, especially the miniprep DNA. It is also very important to inject embryos at the 1 cell stage for gene trap experiments. In our experience, Tol2-mediated transgenesis is very efficient, making it possible to obtain transgenic lines even when injecting later than the 1 cell stage or with lower quality DNA. Substandard injections are much more problematic when carrying out gene trap mutagenesis, since only a small fraction of vector integrations result in effective gene trap events. The second critical step is to ascertain gene trap events by crossing to our UAS:mRFP line. Absence of mRFP expression is almost always indicative of an enhancer trap event. However, sometimes the lack of mRFP expression may indicate silencing of the 14x UAS. It is therefore important to maintain the UAS:mRFP line in a non-silenced state by propagating the next generation using individuals with high RFP expression when crossed to nsftpl6.
We can foresee making several improvements to our gene trap vectors and protocol. The first is to use a less repetitive UAS in the gene trap construct. It has been shown that a 5x UAS is sufficient to achieve full activation in cell culture experiments, and that less repetitive UAS sequences are less susceptible to methylation and silencing 6,25. It may also be possible to design gene trap vectors for fully conditional mutagenesis, in analogy to the vectors used by the international mouse gene trap consortium and recently adapted for zebrafish 2,26,27. Another improvement would be to use a different transposon system, for example Sleeping Beauty 28, to generate a UAS:mRFP reporter line. This would enable injection of Tol2-based gene traps directly into the reporter line and screening of F0s by incrossing. Furthermore, this would eliminate the use of two different genetic backgrounds, making the interpretation of loss of function phenotypes more straightforward. Even with these improvements, the general Gal4 gene trap protocol presented here would still be fully applicable.

Insertional mutagenesis has proven to be a powerful approach to dissecting gene function in various model systems from unicellular organisms like bacteria and yeast to multicellular organisms such as mice and plants. Any exogenous DNA (virus, transposon or plasmid) may serve as a mutagen by interrupting essential elements of genes such as exons or promoters. The effective target of such simple approaches is exceedingly small in large complex genomes, since exons comprise only 1-3% of a typical vertebrate genome. Small effective target size can be overcome by very high vector integration rates, as exemplified by the tremendous success of retroviral insertional mutagenesis in zebrafish 11,12. In contrast, introns comprise about 20-30% of vertebrate genomes. In zebrafish, transposon-based insertional mutagenesis vectors which effectively introduce null- or severe hypomorphic mutations upon integration into introns have been named "gene breaking transposons" (GBTs) 8-10. For efficient gene trap integration, they use a minimal Tol2 transposon 13,14. Mutagenicity of GBTs relies on fish-derived splice acceptor and transcriptional termination/polyadenylation sequences. The elements responsible for GBT mutagenicity are flanked by direct loxP sites for excision by Cre recombinase. Thus, injection of Cre mRNA leads to efficient reversion of GBT-induced mutations, even though some transposon sequences remain at the integration locus 9,10.
The recently published GBT vectors use mRFP as the gene trap reporter 9,10, leading to two potential shortcomings. First, it is not known what fraction of zebrafish genes are expressed at a high enough level to be detected by direct fluorescent reporter fusion proteins. Second, only a small subset of genes are expected to have essential functions. It has been estimated that there are only 1,400-2,400 genes required for zebrafish development 15,16. Most gene trap mutants are not expected to display overt phenotypes and therefore will have limited utility. To overcome these two limitations, we have modified GBT-R15 9,10 to use with Gal4-VP16 as the primary gene trap reporter (Balciuniene et al. in preparation). As with AUG-less mRFP in GBT-R15, the translation start site was removed from Gal4-VP16. For direct detection of gene trap events, our vectors contain an eGFP reporter under the control of 14x Gal4 UAS 17,18.
Several additional features are engineered into our bipartite gene trap vector. The UAS:eGFP cassette is flanked by direct FRT sites (grey chevrons in Figure 1). Injection of Flp recombinase mRNA leads to excision of the UAS:eGFP cassette, leaving the gene trap mutation unmarked by eGFP fluorescence. It can then be used in transgenic lines which mark specific tissues or developmental events by GFP fluorescence. As in other GBTs, the whole gene trap cassette is flanked by direct loxP sites (open pentagons in Figure 1). This makes gene trap events reversible by expression of Cre recombinase, readily establishing proof of causality relationship between a specific gene trap integration and observed phenotype (Figure 2). Finally, the gene trap cassette is flanked by inverted I-SceI meganuclease sites (black triangles in Figure 1). In Drosophila, transposon integrations are often converted to deletions (deficiencies) by imprecise excision of the P element. There is no evidence that excision of Tol2 transposon (or any other transposon active in vertebrates such as Sleeping Beauty, PiggyBac or Ac/Ds) can lead to deletions. We therefore included I-SceI sites as a potential surrogate method for induction of deletions, even though there is no evidence that I-SceI can induce deletions in zebrafish. It should be noted that while I-SceI meganuclease can be used to facilitate transgenesis in zebrafish, DNA cleavage by I-SceI meganuclease is used to study DNA repair mechanisms, including error-prone non-homologous end joining, in yeast and mammalian cells 19-22.
Integration of our gene trap vector can lead to eGFP expression by two different mechanisms. The first is a true gene trap event (Figure 1C): the vector integrates into a gene (IMG for Insertionally Mutated Gene), a fusion transcript between the 5' of the endogenous IMG transcript and the Gal4-VP16 is made and translated into a fusion protein containing the N-terminus of the protein encoded by IMG and Gal4-VP16. This fusion protein binds to the 14x UAS and activates transcription of eGFP. The second, less desirable event is an enhancer trap (Figure 1D): the minimal promoter in front of eGFP falls under the control of an enhancer near the integration site, leading to production of eGFP in the absence of Gal4-VP16 production. In our estimate, 30-50% of eGFP expression events are due to an enhancer trap and 50-70% are due to a gene trap 23(Balciuniene et al. in preparation). To distinguish between these two classes of events, we have made a 14xUAS:mRFP transgenic line (Figure 1B), for convenience marked by lens-specific &gamma;Cry:GFP (Balciuniene et al. in preparation). Gene trap events are verified by co-expression of GFP and RFP (compare C and D in Figure 2).
In our pilot screen, we recovered over 40 gene trap events after screening 270 F0 fish injected with a mixture of transposon DNA and transposase mRNA. Two of our gene trap integrations have occurred into genes with previously published chemically-induced mutants: nsf and flr. The phenotypes of our insertional mutants nsftpl6 and flrtpl24 are superficially indistinguishable from the phenotypes of the corresponding chemically-induced mutants. We also noted that gene trap events appear biased toward the introns near the 5' end of genes, thus increasing the probability of null alleles. We do observe some degree of UAS silencing (variegation) in all of our gene trap lines, and some loci are clearly more susceptible to variegation than others. We do not believe that UAS silencing is a significant problem for propagation of gene trap lines, as we have been able to easily propagate all of our gene trap lines through at least five generations by selecting for GFP expression alone.

<table><tbody><tr><td>Miniprep Kit</td><td>Qiagen</td><td>27104</td><td>Optional PB wash step is a must</td></tr><tr><td>XbaI restriction enzyme</td><td>ThermoFisher</td><td>ER0681</td><td></td></tr><tr><td>mMessage Machine T3</td><td>Ambion</td><td>AM1348</td><td></td></tr><tr><td>RiboLock RNase Inhibitor</td><td>ThermoFisher</td><td>EO0381</td><td></td></tr><tr><td>RNeasy MinElute</td><td>Qiagen</td><td>74204</td><td>Can be replaced by phenol extraction and precipitation. Do not use LiCl!</td></tr><tr><td>Nuclease-free water</td><td>Ambion</td><td>AM9937</td><td>Has to be non-DEPC treated</td></tr><tr><td>Borosilicate glass capiliaries for microinjection needles</td><td>World Precision Instruments</td><td>1B100F-4</td><td></td></tr><tr><td>Microcapiliaries for calibration</td><td>Drummond Scientific</td><td>1-000-0010</td><td></td></tr><tr><td>Microloader tips</td><td>Eppendorf</td><td>5242 956.003</td><td></td></tr><tr><td>Needle puller *</td><td>Sutter Instruments</td><td>P-87</td><td></td></tr><tr><td>Stereomicroscope for microinjection * </td><td>Nikon</td><td>SMZ1500</td><td>We also use Wild 5M and Olympus SZ61 with transmitted light stand</td></tr><tr><td>Picoinjector *</td><td>Harvard Instruments</td><td>Pli-90</td><td>We also use Pli-100</td></tr><tr><td>Screening microscope with GFP/RFP fluorescence *</td><td>Zeiss</td><td>Zeiss AxioImager</td><td>5X Fluar objective has sufficient working distance</td></tr><tr><td>Screening microscope with GFP/RFP fluorescence *</td><td>Nikon</td><td>SMZ1500</td><td></td></tr><tr><td colspan="4">* We suggest this equipment only because we successfully use it in our laboratory. In each category, several comparable options from multiple manufacturers are available.</td></tr></tbody></table>

gene trapping using gal4 in zebrafish

Large clutch size and external development of optically transparent embryos make zebrafish an exceptional vertebrate model system for in vivo insertional mutagenesis using fluorescent reporters to tag expression of mutated genes. Several laboratories have constructed and tested enhancer- and gene-trap vectors in zebrafish, using fluorescent proteins, Gal4- and lexA- based transcriptional activators as reporters 1-7. These vectors had two potential drawbacks: suboptimal stringency (e.g. lack of ability to differentiate between enhancer- and gene-trap events) and low mutagenicity (e.g. integrations into genes rarely produced null alleles). Gene Breaking Transposon (GBTs) were developed to address these drawbacks 8-10. We have modified one of the first GBT vectors, GBT-R15, for use with Gal4-VP16 as the primary gene trap reporter and added UAS:eGFP as the secondary reporter for direct detection of gene trap events. Application of Gal4-VP16 as the primary gene trap reporter provides two main advantages. First, it increases sensitivity for genes expressed at low expression levels. Second, it enables researchers to use gene trap lines as Gal4 drivers to direct expression of other transgenes in very specific tissues. This is especially pertinent for genes with non-essential or redundant functions, where gene trap integration may not result in overt phenotypes. The disadvantage of using Gal4-VP16 as the primary gene trap reporter is that genes coding for proteins with N-terminal signal sequences are not amenable to trapping, as the resulting Gal4-VP16 fusion proteins are unlikely to be able to enter the nucleus and activate transcription. Importantly, the use of Gal4-VP16 does not pre-select for nuclear proteins: we recovered gene trap mutations in genes encoding proteins which function in the nucleus, the cytoplasm and the plasma membrane.

In a successful injection, at least 80% of the embryos injected with the gene trap DNA/ Tol2 transposase mRNA mix display some degree of GFP fluorescence (Figure 3). When the brightest GFP-positive embryos (Figure 3C) are selected to establish the F0 pool, about one in 10 screened F0 fish will yield gene trap progeny. Among the F0 fish from which gene trap events are recovered, most transmit a single gene trap event in addition to several non-expressing transposon integrations. However, we have established up to four gene trap lines from a single F0 individual. GFP/RFP expression patterns in gene trap lines range from extremely tissue specific (for example in olfactory bulbs) to fairly ubiquitous.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50113/50113fig1highres.jpg" src="/files/ftp_upload/50113/50113fig1.jpg" /> 
Figure 1. The Gal4-containing bipartite gene trap vector GBT-B1 and its applications. A. Components of the Gal4-containing gene trap vector: Tol2 5' and Tol2 3', minimal Tol2 transposon ends 13; cSA, Carp &beta;-Actin splice acceptor 8; ^Gal4-VP16, AUG-less Gal4-VP16; zp(A), zebrafish &beta;-Actin 3' UTR and poly(A) elements from GBT-R15 9,10. 14XUAS, eGFP and SV40 p(A) are components of UAS:eGFP expression cassette 17,18. The arrow indicates activation of UAS:eGFP by Gal4-VP16 produced by the gene trap. B. The 14XUAS:mRFP transgene marked by a lens-specific &gamma;Cry:GFP cassette. The arrow indicates activation of UAS:mRFP by Gal4-VP16 in trans. C. Gene trap event. Blue boxes are exons. Splicing is indicated by dashed line. D. Enhancer trap event. Integration of the vector near an enhancer (brown box) may place the minimal eGFP promoter under the control of this enhancer (brown arrow). This would result in a tissue-specific expression of eGFP, but since Gal4-VP16 is not made, mRFP expression would not be activated.
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/50113/50113fig2highres.jpg" src="/files/ftp_upload/50113/50113fig2.jpg" /> 
Figure 2. Reversion of gene trap events using Cre recombinase. A, B. Diagram of a gene trap mutation (A) and Cre-reverted gene trap allele (B). C, D. Co-expression of GFP (C) and RFP (D) in the nervous system of the nsftpl6 gene trap line. E, F. Injection of Cre RNA into nsftpl6 embryos leads to loss of GFP (E) and RFP (F) expression. Note that there is no reduction of GFP expression in the lens, as expected.
<img alt="Figure 3" src="/files/ftp_upload/50113/50113fig3.jpg" /> 
Figure 3. Embryos injected with the GBT-B1 (pDB783) gene trap. A. Embryos injected with pGBT-B1 (pDB783) alone display little GFP fluorescence in the body. B, C. Embryos injected with pGBT-B1 and Tol2 transposase mRNA: a random group of embryos (B) and high-expressors selected for rearing (C).
<img alt="Figure 4" fo:content-width="6in" fo:src="/files/ftp_upload/50113/50113fig4highres.jpg" src="/files/ftp_upload/50113/50113fig4.jpg" /> 
Figure 4. Experimental outline for establishment of Gal4 gene trap lines. Partly overlapping red/green structures indicate co-expression of GFP and RFP, while green alone indicates mosaicism for gene trap or enhancer trap events.

Watch this Scientific Journal Video about Gene Trapping Using Gal4 in Zebrafish at JoVE.com

Gene Trapping Using Gal4 in Zebrafish

This protocol describes the method of gene trap insertional mutagenesis using Gal4-VP16 as the primary reporter and GFP/RFP as secondary reporters in zebrafish. Approximately one in ten high-expressing F0 fish yield gene trap progeny co-expressing GFP and RFP. The screening procedure can be readily scaled to adapt to the size of the laboratory performing the insertional mutagenesis screen.

Large clutch size and external development of optically transparent embryos make zebrafish an exceptional vertebrate model system for in vivo insertional ...

Research

JoVE Journal

Biology

18.2K Views.  Temple University. This protocol describes the method of gene trap insertional mutagenesis using Gal4-VP16 as the primary reporter and GFP/RFP as secondary reporters in zebrafish. Approximately one in ten high-expressing F0 fish yield gene trap progeny co-expressing GFP and RFP. The screening procedure can be readily scaled to adapt to the size of the laboratory performing the insertional mutagenesis screen.

Video: Gene Trapping Using Gal4 in Zebrafish

Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS expression system1 in zebrafish larvae, and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells. The KalTA4-ERT2/UAS system is both inducible and reversible which allows us to induce cell transformation with precise temporal/spatial resolution in vivo. This provides us with a unique opportunity to live image how individual preneoplastic cells interact with host tissues as soon as they emerge, then follow their progression as well as regression. Recent studies in zebrafish larvae have shown a trophic function of innate immunity in the earliest stages of tumorigenesis2,3. Our inducible system would allow us to live image the onset of cellular transformation and the subsequent host response, which may lead to important insights on the underlying mechanisms for the regulation of oncogenic trophic inflammatory responses. We also discuss how one might adapt our protocol to achieve temporal and spatial control of ectopic gene expression in any tissue of interest.

Studying the earliest events of preneoplastic cell progression and innate immune cell interaction is pivotal to understand and treat cancer. Here we describe a method to conditionally induce epithelial cell transformations and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells in zebrafish larvae.

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS ...

Developmental Biology

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos. Recent advances in gene editing tools are enabling precise introduction of mutations and transgenes in zebrafish. Disease modeling in zebrafish often leads to larval phenotypes and early death which can be challenging to interpret if genotypes are unknown. This early identification of genotypes is also needed in experiments requiring sample pooling, such as in gene expression or mass spectrometry studies. However, extensive genotypic screening is limited by traditional methods, which in most labs are performed only on adult zebrafish or in postmortem larvae. We addressed this problem by adapting a method for the isolation of PCR-ready genomic DNA from live zebrafish larvae that can be achieved as early as 72 h post-fertilization (hpf). This time and cost-effective technique, improved from a previously published genotyping protocol, allows the identification of genotypes from microscopic fin biopsies. The fins quickly regenerate as the larvae develop. Researchers are then able to select and raise the desired genotypes to adulthood by utilizing this high-throughput PCR-based genotyping procedure.

Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short ...

Genetics

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific regions of developing embryos, including those of C. elegans, chick, Xenopus, zebrafish, and mouse 1. In zebrafish, in vivo electroporation has been shown to have excellent spatial and temporal resolution for the delivery of these reagents 2-7. The temporal resolution of this method is important because it allows for incorporation of these reagents at specific stages in development. Furthermore, because expression from electroporated vectors occurs within 6 hours 7, this method is more timely than transgenic approaches. While the spatial resolution can be extremely precise when targeting a single cell 2, 6, it is often preferable to incorporate reagents into a specific cell population within a tissue or structure. When targeting multiple cells, in vivo electroporation is efficient for delivery to a specific region of the embryo; however, particularly within the developing nervous system, it is difficult to target specific cell types solely through spatially discrete electroporation. Alternatively, enhancer trap transgenic lines offer excellent cell type-specific expression of transgenes 8. Here we describe an approach that combines transgenic Gal4-based enhancer trap lines 8 with spatially discrete in vivo electroporation 7, 9 to specifically target developing neurons of the zebrafish olfactory bulb. The Et(zic4:Gal4TA4,UAS:mCherry)hzm5 (formerly GA80_9) enhancer trap line previously described 8, displays targeted transgenic expression of mCherry mediated by a zebrafish optimized Gal4 (KalTA4) transcriptional activator in multiple regions of the developing brain including hindbrain, cerebellum, forebrain, and the olfactory bulb. To target GFP expression specifically to the olfactory bulb, a plasmid with the coding sequence of GFP under control of multiple Gal4 binding sites (UAS) was electroporated into the anterior end of the forebrain at 24-28 hours post-fertilization (hpf). Although this method incorporates plasmid DNA into multiple regions of the forebrain, GFP expression is only induced in cells transgenically expressing the KalTA4 transcription factor. Thus, by using the GA080_9 transgenic line, this approach led to GFP expression exclusively in the developing olfactory bulb. GFP expressing cells targeted through this approach showed typical axonal projections, as previously described for mitral cells of the olfactory bulb 10. This method could also be used for targeted delivery of other reagents including short-hairpin RNA interference expression plasmids, which would provide a method for spatially and temporally discrete loss-of-function analysis.

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

The temporal and spatial resolution of genetic manipulations determines the spectrum of biological phenomena that they can perturb. Here we use temporally and spatially discrete in vivo electroporation, combined with transgenic lines of zebrafish, to induce expression of a GFP transgene specifically in neurons of the developing olfactory bulb.

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific ...

Neuroscience

Manipulation of Gene Function in Mexican Cavefish

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, including eye degeneration, albinism, sleep loss, hyperphagia, and sensory processing. Species of cavefish from around the world display a convergent evolution of morphological and behavioral traits due to shared environmental pressures between different cave systems. Diverse cave species have been studied in the laboratory setting. The Mexican tetra, Astyanax mexicanus, with sighted and blind forms, has provided unique insights into biological and molecular processes underlying the evolution of complex traits and is well-poised as an emerging model system. While candidate genes regulating the evolution of diverse biological processes have been identified in A. mexicanus, the ability to validate a role for individual genes has been limited. The application of transgenesis and gene-editing technology has the potential to overcome this significant impediment and to investigate the mechanisms underlying the evolution of complex traits. Here, we describe a different methodology for manipulating gene expression in A. mexicanus. Approaches include the use of morpholinos, Tol2 transgenesis, and gene-editing systems, commonly used in zebrafish and other fish models, to manipulate gene function in A. mexicanus. These protocols include detailed descriptions of timed breeding procedures, the collection of fertilized eggs, injections, and the selection of genetically modified animals. These methodological approaches will allow for the investigation of the genetic and neural mechanisms underlying the evolution of diverse traits in A. mexicanus.&#160;

We describe approaches for the manipulation of genes in the evolutionary model system Astyanax mexicanus. Three different techniques are described: Tol2-mediated transgenesis, targeted manipulation of the genome using CRISPR/Cas9, and knockdown of expression using morpholinos. These tools should facilitate the direct investigation of genes underlying the variation between surface- and cave-dwelling forms.

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, ...

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression studies. In zebrafish (Danio rerio), microinjection of RNA, DNA, proteins, antisense oligonucleotides and other small molecules into the developing embryo provides researchers a quick and robust assay for exploring gene function in vivo. In this video-article, we will demonstrate how to prepare and microinject in vitro synthesized EGFP mRNA and a translational-blocking morpholino oligo against pkd2, a gene associated with autosomal dominant polycystic kidney disease (ADPKD), into 1-cell stage zebrafish embryos. We will then analyze the success of the mRNA and morpholino microinjections by verifying GFP expression and phenotype analysis. Broad applications of this technique include generating transgenic animals and germ-line chimeras, cell-fate mapping and gene screening. Herein we describe a protocol for overexpression of EGFP and knockdown of pkd2 by mRNA and morpholino oligonucleotide injection.

Microinjection is a well-established and effective method for introducing foreign substances into fertilized zebrafish embryos. Here, we demonstrate a robust microinjection technique for performing mRNA overexpression, and morpholino oligonucleotide gene knockdown studies in zebrafish.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Fluorescent Protein Tagging in Zebrafish Larvae Using CRISPR/Cas9 Knock-In

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in vivo imaging due to its transparency during the early developmental stage. Here, we provide a detailed protocol for performing high-efficiency fluorescence gene KI, rapid screening for KI founders, and low-abundance protein tracing in zebrafish larvae, which will lay a critical foundation for subsequent physio-pathological studies in zebrafish. The current protocol includes complete steps for the sgRNA design for the gene of interest, sgRNA in vitro transcription, Cas9 mRNA in vitro transcription, in vivo sgRNA screen for the one with the highest efficiency, donor plasmid design and construction, microinjection in zebrafish larvae, KI founder screen and zebrafish live imaging. Critical steps, troubleshooting tips, quality control methods, and advantages and applications of this protocol are included and discussed. This protocol assures quick and accurate results at a low cost and has been validated by multiple trials.

Here, we present a protocol to label proteins with a fluorescence protein tag in zebrafish larvae, a newly developed and modified in vivo system especially useful for visualizing low-abundance proteins in zebrafish.

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in ...

The Zebrafish Tol2 System: A Modular and Flexible Gateway-Based Transgenesis Approach

Fetal alcohol spectrum disorders (FASD) are characterized by a highly variable set of structural defects and cognitive impairments that arise due to prenatal ethanol exposure. Due to the complex pathology of FASD, animal models have proven critical to our current understanding of ethanol-induced developmental defects. Zebrafish have proven to be a powerful model to examine ethanol-induced developmental defects due to the high degree of conservation of both genetics and development between zebrafish and humans. As a model system, zebrafish possess many attributes that make them ideal for developmental studies, including large numbers of externally fertilized embryos that are genetically tractable and translucent. This allows researchers to precisely control the timing and dosage of ethanol exposure in multiple genetic contexts. One important genetic tool available in zebrafish is transgenesis. However, generating transgenic constructs and establishing transgenic lines can be complex and difficult. To address this issue, zebrafish researchers have established the transposon-based Tol2 transgenesis system. This modular system uses a multisite Gateway cloning approach for the quick assembly of complete Tol2 transposon-based transgenic constructs. Here, we describe the flexible Tol2 system toolbox and a protocol for generating transgenic constructs ready for zebrafish transgenesis and their use in ethanol studies.

This work describes a protocol for the modular Tol2 transgenesis system, a gateway-based cloning method to create and inject transgenic constructs into zebrafish embryos.

Fetal alcohol spectrum disorders (FASD) are characterized by a highly variable set of structural defects and cognitive impairments that arise due to ...

Gene Trapping Using Gal4 in Zebrafish

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Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Imaging Subcellular Structures in the Living Zebrafish Embryo

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

Manipulation of Gene Function in Mexican Cavefish

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

The Zebrafish Tol2 System: A Modular and Flexible Gateway-Based Transgenesis Approach