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 border="1">
<tbody>
 <tr>
 <td>Name of the Reagent/Material</td>
 <td>Company</td>
 <td>Catalog #</td>
 <td>Comments</td>
 </tr>
 <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></td>
 <td></td>
 <td></td>
 <td>* 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 ...

Universidad de Cartagena UDEC

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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

Microinjection of Zebrafish Embryos to Analyze Gene Function

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

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

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

Global Gene Expression Analysis Using a Zebrafish Oligonucleotide Microarray Platform

Gene microarray technology permits quantitative measurement and gene expression profiling of transcript levels on a genome-wide basis. Gene microarray technology is used in numerous biological disciplines in a variety of applications including global gene expression analysis in relation to developmental stage, to a disease state, and in toxic responses. Herein, we include a demonstration of global gene expression analysis using a comprehensive zebrafish-specific oligonucleotide microarray platform. The zebrafish expression microarray platform contains 385,000 probes, 60 base pairs in length, interrogating 37,157 targets with up to 12 probes per target. For this platform, all cDNA and genomic information available for the zebrafish was collected from various genomic databases including Ensembl (http://www.ensembl.org), VEGA (http://vega.sanger.ac.uk), UCSC (http://genome.ucsc.edu), and ZFIN (http://www.zfin.org). As a result this expression array provides complete coverage of the current zebrafish transcriptome. The zebrafish expression microarray was printed by Roche NimbleGen (Madison, WI). This technical demonstration includes the fluorescent labeling of a cDNA product, hybridization of the labeled cDNA product to the microarray platform, and array scanning for signal acquisition using the one color analysis strategy.  

Gene microarrays are powerful tools in gene expression profiling at a genome-wide level. This technology has application in a variety of biological disciplines including developmental biology and toxicology. In this video, we detail a protocol for global gene expression analysis using a comprehensive oligonucleotide microarray platform for the zebrafish.

Gene microarray technology permits quantitative measurement and gene expression profiling of transcript levels on a genome-wide basis. Gene microarray ...

Using the Gene Pulser MXcell Electroporation System to Transfect Primary Cells with High Efficiency

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser MXcell electroporation system and Gene Pulser electroporation buffer (Bio-Rad) were specifically developed to easily transfect nucleic acids into mammalian cells and difficult-to-transfect cells, such as primary and stem cells. We will demonstrate how to perform a simple experiment to quickly identify the best electroporation conditions. We will demonstrate how to run several samples through a range of electroporation conditions so that an experiment can be conducted at the same time as optimization is performed. We will also show how optimal conditions identified using 96-well electroporation plates can be used with standard electroporation cuvettes, facilitating the switch from electroporation plates to electroporation cuvettes while maintaining the same electroporation efficiency. In the video, we will also discuss some of the key factors that can lead to the success or failure of electroporation experiments.

This procedure shows how to use the Gene Pulser MXcell electroporation system to rapidly and easily identify the best electroporation conditions for mouse embryonic fibroblasts (MEFs) or other primary cells. Considerations for troubleshooting are also discussed in the associated video.

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser ...

Using an Automated Cell Counter to Simplify Gene Expression Studies: siRNA Knockdown of IL-4 Dependent Gene Expression in Namalwa Cells

The use of siRNA mediated gene knockdown is continuing to be an important tool in studies of gene expression.  siRNA studies are being conducted not only to study the effects of downregulating single genes, but also to interrogate signaling pathways and other complex interaction networks.  These pathway analyses require both the use of relevant cellular models and methods that cause less perturbation to the cellular physiology.  Electroporation is increasingly being used as an effective way to introduce siRNA and other nucleic acids into difficult to transfect cell lines and primary cells without altering the signaling pathway under investigation. There are multiple critical steps to a successful siRNA experiment, and there are ways to simplify the work while improving the data quality at several experimental stages.  To help you get started with your siRNA mediated gene knockdown project, we will demonstrate how to perform a pathway study complete from collecting and counting the cells prior to electroporation through post transfection real-time PCR gene expression analysis.  The following study investigates the role of the transcriptional activator STAT6 in IL-4 dependent gene expression of CCL17 in a Burkitt lymphoma cell line (Namalwa).  The techniques demonstrated are useful for a wide range of siRNA-based experiments on both adherent and suspension cells.  We will also show how to streamline cell counting with the TC10 automated cell counter, how to electroporate multiple samples simultaneously using the MXcell electroporation system, and how to simultaneously assess RNA quality and quantity with the Experion automated electrophoresis system.

This procedure describes a quick and easy workflow to introduce siRNA into difficult to transfect cell lines and follow gene expression by real-time PCR. Use of an automated cell counter, multi-well electroporation plate, and automated electrophoresis station provide quick and reliable results without the need for expensive robotic handling.

The use of siRNA mediated gene knockdown is continuing to be an important tool in studies of gene expression.  siRNA studies are being conducted not only ...

Induction and Testing of Hypoxia in Cell Culture

Hypoxia is defined as the reduction or lack of oxygen in organs, tissues, or cells. This decrease of oxygen tension can be due to a reduced supply in oxygen (causes include insufficient blood vessel network, defective blood vessel, and anemia) or to an increased consumption of oxygen relative to the supply (caused by a sudden higher cell proliferation rate). Hypoxia can be physiologic or pathologic such as in solid cancers 1-3, rheumatoid arthritis, atherosclerosis etc&#x2026; Each tissues and cells have a different ability to adapt to this new condition. During hypoxia, hypoxia inducible factor alpha (HIF) is stabilized and regulates various genes such as those involved in angiogenesis or transport of oxygen 4. The stabilization of this protein is a hallmark of hypoxia, therefore detecting HIF is routinely used to screen for hypoxia 5-7. 
In this article, we propose two simple methods to induce hypoxia in mammalian cell cultures and simple tests to evaluate the hypoxic status of these cells.

Here we propose simple methods to induce hypoxia in cell cultures and simple tests to evaluate the hypoxic status of the cultures.

Hypoxia is defined as the reduction or lack of oxygen in organs, tissues, or cells.  This decrease of oxygen tension can be due to a reduced supply in ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the messagefor review see 1. However, it's now widely accepted that synonymous codon usage is the primary cause of non-uniform translational elongation rates1. Synonymous codons are not used with identical frequency. A bias exists in the use of synonymous codons with some codons used more frequently than others2. Codon bias is organism as well as tissue specific2,3. Moreover, frequency of codon usage is directly proportional to the concentrations of cognate tRNAs4. Thus, a frequently used codon will have higher multitude of corresponding tRNAs, which further implies that a frequent codon will be translated faster than an infrequent one. Thus, regions on mRNA enriched in rare codons (potential pause sites) will as a rule slow down ribosome movement on the message and cause accumulation of nascent peptides of the respective sizes5-8. These pause sites can have functional impact on the protein expression, mRNA stability and protein foldingfor review see 9. Indeed, it was shown that alleviation of such pause sites can alter ribosome movement on mRNA and subsequently may affect the efficiency of co-translational (in vivo) protein folding1,7,10,11. To understand the process of protein folding in vivo, in the cell, that is ultimately coupled to the process of protein synthesis it is essential to gain comprehensive insights into the impact of codon usage/tRNA content on the movement of ribosomes along mRNA during translational elongation.
Here we describe a simple technique that can be used to locate major translation pause sites for a given mRNA translated in various cell-free systems6-8. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA. The rationale is that at low-frequency codons, the increase in the residence time of the ribosomes results in increased amounts of nascent peptides of the corresponding sizes. In vitro transcribed mRNA is used for in vitro translational reactions in the presence of radioactively labeled amino acids to allow the detection of the nascent chains. In order to isolate ribosome bound nascent polypeptide complexes the translation reaction is layered on top of 30% glycerol solution followed by centrifugation. Nascent polypeptides in polysomal pellet are further treated with ribonuclease A and resolved by SDS PAGE. This technique can be potentially used for any protein and allows analysis of ribosome movement along mRNA and the detection of the major pause sites. Additionally, this protocol can be adapted to study factors and conditions that can alter ribosome movement and thus potentially can also alter the function/conformation of the protein.

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

A technique to identify translational pause sites on mRNA is described. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA, followed by the size analysis of the nascent chains using a denaturing gel electrophoresis.

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the ...

Using SecM Arrest Sequence as a Tool to Isolate Ribosome Bound Polypeptides

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway that polypeptide chain follows during co-translational folding to achieve its functional form is still an enigma. In order to understand this process and to determine the exact conformation of the co-translational folding intermediates, it is essential to develop techniques that allow the isolation of RNCs carrying nascent chains of predetermined sizes to allow their further structural analysis.
SecM (secretion monitor) is a 170 amino acid E. coli protein that regulates expression of the downstream SecA (secretion driving) ATPase in the secM-secA operon6. Nakatogawa and Ito originally found that a 17 amino acid long sequence (150-FSTPVWISQAQGIRAGP-166) in the C-terminal region of the SecM protein is sufficient and necessary to cause stalling of SecM elongation at Gly165, thereby producing peptidyl-glycyl-tRNA stably bound to the ribosomal P-site7-9. More importantly, it was found that this 17 amino acid long sequence can be fused to the C-terminus of virtually any full-length and/or truncated protein thus allowing the production of RNCs carrying nascent chains of predetermined sizes7. Thus, when fused or inserted into the target protein, SecM stalling sequence produces arrest of the polypeptide chain elongation and generates stable RNCs both in vivo in E. coli cells and in vitro in a cell-free system. Sucrose gradient centrifugation is further utilized to isolate RNCs.
The isolated RNCs can be used to analyze structural and functional features of the co-translational folding intermediates. Recently, this technique has been successfully used to gain insights into the structure of several ribosome bound nascent chains10,11. Here we describe the isolation of bovine Gamma-B Crystallin RNCs fused to SecM and generated in an in vitro translation system.

We describe here a technique that is now routinely used to isolate stably bound ribosome nascent chain complexes (RNCs). This technique takes advantage of the discovery that a 17 amino acid long SecM "arrest sequence" can halt translation elongation in a prokaryotic (E. coli) system, when inserted into (or fused to the C-terminus) of virtually any protein.

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway ...

Gavaging Adult Zebrafish

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or agents in solutions for scientific research. Current methods for administering compounds orally to adult zebrafish are inaccurate due to variability in voluntary consumption by the fish. A gavage procedure was developed to deliver precise quantities of infectious agents to zebrafish for study in biomedical research. Adult zebrafish over 6 months of age were anesthetized with 150 mg/L of buffered MS-222 and gavaged with 5 &mu;l of solution using flexible catheter implantation tubing attached to a cut 22-G needle tip. The flexible tubing was lowered into the oral cavity of the zebrafish until the tip of the tubing extended past the gills (approximately 1 cm). The solution was then injected slowly into the intestinal tract. This method was effective 88% of the time, with fish recovering uneventfully. This procedure is also efficient as one person can gavage 20-30 fish in one hour. This method can be used to precisely administer agents for infectious diseases studies, or studies of other compounds in adult zebrafish.

The increasing use of zebrafish as an animal model requires the development of effective methods for the delivery of known quantities of compounds and solutions. The gavage procedure described below allows for the oral delivery of precise volumes of solution reliably, safely and efficiently to adult zebrafish.

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Gene Trapping Using Gal4 in Zebrafish

Summary

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Chapters in this video

Microinjection of Zebrafish Embryos to Analyze Gene Function

Global Gene Expression Analysis Using a Zebrafish Oligonucleotide Microarray Platform

Using the Gene Pulser MXcell Electroporation System to Transfect Primary Cells with High Efficiency

Using an Automated Cell Counter to Simplify Gene Expression Studies: siRNA Knockdown of IL-4 Dependent Gene Expression in Namalwa Cells

Induction and Testing of Hypoxia in Cell Culture

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

Using SecM Arrest Sequence as a Tool to Isolate Ribosome Bound Polypeptides

Gavaging Adult Zebrafish

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy