This work was funded in part by NIH R01 #DE021475 (CTM), an NIH Predoctoral Training Grant 5T32GM007127 (PAE), and an NSF Graduate Research Fellowship (NAE). We thank Kevin Schwalbach for performing BAC recombineering and injections, Nick Donde for generating CRISPR Sanger sequencing data, and Katherine Lipari for helpful feedback on the injection protocol.

The authors have nothing to disclose.

Injecting one-cell stickleback embryos for transgenesis or genome editing presents three main challenges. First, relative to zebrafish embryos, the stickleback embryonic chorion is tough and will often break needles. This problem can be partially overcome by using thicker and stronger glass micropipettes and injecting perpendicular to the chorion (see Protocol, Figure 2). Ensuring that as little water as possible is added to the embryos (just enough to cause the chorion to swell and lift away from the cell) helps to reduce chorion hardness. The chorion hardens over time, so working quickly after moistening the embryos is important. Keeping the embryos in a humidity chamber prior to injection so that they do not dry out is also critical. Some clutches and even individual embryos simply have much thicker and tougher chorions; sometimes moving on to the next embryo or trying with a new clutch is the easiest solution. It is much easier to skip one difficult embryo than to replace a damaged needle. Having backup needles ready will allow injections to continue in the case of needle breakage. When injecting BACs, it is common for the needle to clog. The needle can be unclogged by gently scraping or re-breaking the tip with forceps, or by using the constant air pressure switch to purge the clog.
Second, identifying the first cell in the embryo is challenging; it is often quite flattened and difficult for beginners to see, and is especially invisible when looking directly at it. Often the blastomere can only be seen as a slightly raised bump in profile. Therefore, it is best to identify the cell in profile (Figure 3B) and then gently rotate the embryo forward and to the side so the cell will face the end of the angled needle (though the cell will often be invisible from this angle; the darker color of the blastomere in Figure 3 is exaggerated for clarity).
Third, targeting the cytoplasm is also difficult, especially if the first cell is especially flat. Aiming for the fattest part of the blastomere (usually the center) improves the chance of injecting into cytoplasm. While injecting into the yolk near the cytoplasm can successfully produce transgenic fish, the efficiency seems to be increased and lethality decreased when the cytoplasm is targeted with a single needle puncture. Sometimes, individual clutches will have particularly thin blastomeres, such that avoiding the yolk is nearly impossible. Waiting longer than 25 min to begin injections may help (some clutches do not form a full size blastomere until ~45 min post fertilization), but if the blastomeres never increase in size, it is often easier to obtain a new clutch of eggs than to try to inject flattened cells.
Excessive lethality following injections may occur for several reasons. Blunt needles and/or too large a needle bore size may cause too much damage to the cell and/or cause cytoplasm to leak out. Some DNA constructs seem to be especially lethal; lowering the concentration of DNA may improve survival but lower transgenesis rates. Cleaning up plasmids first with a midiprep kit that contains an endotoxin rinse followed by a second PCR cleanup kit reduces construct toxicity. Finally, genome editing may produce a loss of function mutation that is lethal to developing embryos. Reducing the concentration of the CRISPR or TALEN mRNAs can increase the mosaicism of the embryo to prevent lethality, but may reduce mutant allele recovery efficiency.
A previously published protocol for generating transgenic sticklebacks using a meganuclease method21 reported a 4-7% transgene germline transmission rate from F0 founder fish. The Tol2 method reported here resulted in up to a 72% transgene germline transmission rate from F0 founder fish (indicating multiple genomic integrations). The previous study using a meganuclease method reported 40% of injected embryos showing specific GFP expression, similar to that reported here. Thus while similar rates of transgenic F0 founders are observed for transgenic fish generated by both the meganuclease and Tol2 methods, the germline transmission rate appears much higher for Tol2 mediated transgenics.
As genome editing technologies continue to advance, even more specific genetic manipulations will become possible in stickleback and other fish species. For example, directed repair46 and homologous recombination will allow allele swaps between marine and freshwater stickleback genomes, and modified CRISPRs can be used to specifically activate or inhibit gene expression47. These exciting technologies for genome editing and analysis will lead to new insights about the genetic basis of many interesting morphological, physiological, and behavioral phenotypes in sticklebacks and other fish species.

One fundamental component of understanding how biodiversity arises is determining the genetic and developmental bases of evolved phenotypic changes in nature. The threespine stickleback fish, Gasterosteus aculeatus, has emerged as an excellent model for studying the genetic basis of evolution. Sticklebacks have undergone many adaptive evolutionary changes as marine fish have colonized countless freshwater environments around the northern hemisphere, resulting in dramatic morphological, physiological, and behavioral changes1. The genomes of individuals from twenty-one stickleback populations have been sequenced and assembled, and a high density linkage map has been generated to further improve the assembly2,3. Genetic mapping experiments have identified genomic regions underlying evolved phenotypes4-6, and in a few cases, the functional roles of specific candidate genes have been tested7,8. A number of genomic regions underlying morphological changes have been identified with promising candidate genes, but these candidates have not yet been functionally tested9-12. In addition, sticklebacks are common models for studies of population genetics/genomics13,14, speciation15, behavior1, endocrinology16, ecotoxicology17, immunology18 and parasitology19. Future studies in each of these fields will benefit from the ability to perform functional genetic manipulations in sticklebacks. In addition to manipulating their coding sequences, the roles of candidate genes can be assessed by studying their cis-regulatory sequences and by functionally increasing, decreasing, or eliminating expression of the candidate gene. Microinjection and transgenesis methods in sticklebacks are well established7,8,20 and were initially developed using a meganuclease-mediated method21 first described in medaka22. The modified microinjection method presented here has been optimized for both Tol2-mediated transgenesis and recently developed genome editing reagents including TALENs and CRISPRs.
Changes to cis-regulatory elements are thought to be critical to morphological evolution, as cis-regulatory changes can avoid the negative pleiotropic consequences of coding mutations23. Therefore, testing and comparing putative cis-regulatory sequences has become a central goal of an increasing number of evolutionary studies. In addition, most human disease variants are regulatory variants24,25, and model vertebrate systems are sorely needed to study cis-regulatory element function and logic. Fish that fertilize their embryos externally in large numbers offer powerful vertebrate systems to study cis-regulation. The Tol2 transposon system, in which foreign DNA to be integrated in the genome is flanked by Tol2 transposase binding sites and co-injected with Tol2 transposase mRNA, works with high efficiency for successfully integrating plasmid constructs into fish genomes26-28. Typically, a potential enhancer is cloned upstream of a basal promoter (such as hsp70l29) and fluorescent reporter gene such as EGFP (enhanced green fluorescent protein) or mCherry in a Tol2 backbone and injected with transposase mRNA26. Observation of expression of the fluorescent reporter, either in injected embryos or offspring with stably integrated transgenes, provides information about the spatiotemporal regulation of gene expression driven by the putative enhancer. In further experiments, validated enhancers can be used to drive tissue-specific overexpression of genes of interest.
For analysis of larger cis-regulatory regions, high quality large-insert genomic libraries using bacterial artificial chromosomes (BACs) have been constructed for both marine and freshwater sticklebacks30. These BACs can be recombineered to replace a gene with a fluorescent reporter gene in the context of a large (150-200 kb) genomic region31. The fluorescent reporter is then expressed in a spatiotemporal pattern as determined by regulatory sequences within the BAC. For studies in fish, Tol2 sites can be added to the BAC to facilitate genomic integration32,33. In later stages of development when in situ hybridization is technically challenging, the fluorescent readout of the BAC can be used to study patterns of gene expression, as has been shown for stickleback Bone morphogenetic protein 6 (Bmp6)20. Additionally, fluorescent expression patterns in an individual can be tracked over time, which cannot be accomplished with in situ hybridization. BACs can also be used to add an additional copy of a genomic region to increase dosage of a gene of interest.
For the study of gene function, genome editing is an explosively expanding field that can be used to produce targeted changes to genomic sequences in a wide variety of organisms34. Transcription activator-like effector nucleases (TALENs) are modular, sequence-specific nucleases originally isolated from plant pathogens that can be precisely engineered to bind directly to a genomic sequence of choice and generate a double strand break35,36. Clustered regularly interspaced short palindromic repeats (CRISPR)/CAS systems were originally found in bacteria and use a guide RNA and the Cas9&#160;protein to generate a break in a target DNA sequence complementary to the guide37. The subsequent repair of the double strand break created by both TALENs and CRISPRs often leaves behind a small insertion or deletion, which can disrupt the function of the target sequence35-37. In sticklebacks, TALENs have been used to disrupt gene expression by targeting an enhancer20, and both TALENs and CRISPRs have successfully produced mutations in coding sequences (unpublished data). A detailed protocol for the generation of CRISPRs for use in zebrafish can be used as a guideline to develop CRISPRs for sticklebacks38.
Transgenic and genome editing experiments require introduction of nucleic acids into a newly fertilized one-cell embryo. By introducing the transgene or genome-editing tool early in development, the number of genetically manipulated daughter cells in the embryo is maximized. Injected embryos are then visually screened for fluorescence or molecularly screened for genome modifications. If cells contributing to the germline are successfully targeted, the transgene or mutation can be passed on to a subset of offspring, even when post-injection lethality is high. The mosaic fish can be outcrossed or intercrossed and their offspring screened to recover the mutant alleles or a stably integrated transgene of interest. This protocol describes methods for introducing transgenes and genome editing reagents into one-cell stickleback embryos and monitoring for successful genomic modifications.

<table border="0" cellpadding="0" cellspacing="0" width="647">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Stereomicroscope with transillumination</td>
			<td style="width:145px;">Leica</td>
			<td style="width:119px;">S6e/ KL300 LED</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Manual micromanipulator</td>
			<td style="width:145px;">Applied Scientific Instrumentation</td>
			<td style="width:119px;">MM33</td>
			<td style="width:167px;">Marzhauser M33 Micromanipulator</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Pressure Injecion system</td>
			<td style="width:145px;">Applied Scientific Instrumentation</td>
			<td style="width:119px;">MPPI-3</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Back pressure unit</td>
			<td style="width:145px;">Applied Scientific Instrumentation</td>
			<td style="width:119px;">BPU</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Micropipette holder kit</td>
			<td style="width:145px;">Applied Scientific Instrumentation</td>
			<td style="width:119px;">MPIP</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Magnetic base holder</td>
			<td style="width:145px;">Applied Scientific Instrumentation</td>
			<td style="width:119px;">Magnetic base</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Foot switch</td>
			<td style="width:145px;">Applied Scientific Instrumentation</td>
			<td style="width:119px;">FSW</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Iron plate (magnetic base)</td>
			<td style="width:145px;">Narishige</td>
			<td style="width:119px;">IP</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Flaming/Brown Micropipette Puller</td>
			<td style="width:145px;">Sutter Instrument</td>
			<td style="width:119px;">P-97</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Disposable transfer pipettes</td>
			<td style="width:145px;">Fisher</td>
			<td style="width:119px;">13-711-7M</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">0.5% phenol red in DPBS</td>
			<td style="width:145px;">Sigma&#160;</td>
			<td style="width:119px;">P0290</td>
			<td style="width:167px;">injection tracer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">#5 forceps, biologie dumoxel&#160;</td>
			<td style="width:145px;">Fine Science Tools</td>
			<td style="width:119px;">11252-30</td>
			<td style="width:167px;">for needle breaking</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Micropipette Storage Jar</td>
			<td style="width:145px;">World Precision Instruments</td>
			<td style="width:119px;">E210</td>
			<td>holds needles</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">6&#34;, 6 teeth per inch plaster drywall saw</td>
			<td style="width:145px;">Lenox</td>
			<td>20571 (S636RP)</td>
			<td>hold eggs for injection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">13 cm x 13 cm glass plate</td>
			<td style="width:145px;">any hardware store</td>
			<td style="width:119px;">-</td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">Borosilicate glass capillaries, 1.0 mm OD/0.58 mm ID&#160;</td>
			<td style="width:145px;">World Precision Instruments</td>
			<td style="width:119px;">1B100-F4</td>
			<td style="width:167px;">*harder glass than zebrafish injection capillaries</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">150 x 15mm petri dish</td>
			<td style="width:145px;">Fisher</td>
			<td style="width:119px;">FB0875714</td>
			<td>raise stickleback embryos</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">35 x 10mm petri dish</td>
			<td style="width:145px;">Fisher</td>
			<td style="width:119px;">08-757-100A</td>
			<td>store eggs pre-injection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Instant Ocean Salt</td>
			<td style="width:145px;">Instant Ocean</td>
			<td>SS15-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium Bicarbonate</td>
			<td style="width:145px;">Sigma</td>
			<td>S5761-500G</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Tricaine methanesulfonate/MS-222</td>
			<td style="width:145px;">Western Chemical Inc</td>
			<td style="width:119px;">MS222</td>
			<td style="width:167px;">fish anaesthesia/euthanasia</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">Sp6 transcription kit</td>
			<td style="width:145px;">Ambion</td>
			<td style="width:119px;">AM1340</td>
			<td style="width:167px;">For transcription of TALENs and transposase mRNA</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">RNeasy cleanup kit</td>
			<td style="width:145px;">Qiagen</td>
			<td style="width:119px;">74104</td>
			<td style="width:167px;">purify transposase or TALEN RNA</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">QiaQuick PCR cleanup kit</td>
			<td style="width:145px;">Qiagen</td>
			<td style="width:119px;">28104</td>
			<td style="width:167px;">clean up plasmids for injection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Proteinase K 20 mg/ml</td>
			<td style="width:145px;">Ambion</td>
			<td style="width:119px;">AM2546</td>
			<td style="width:167px;">for DNA preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Nucleobond BAC 100 kit</td>
			<td style="width:145px;">Clontech</td>
			<td style="width:119px;">740579</td>
			<td>for BAC DNA preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">NotI</td>
			<td style="width:145px;">NEB</td>
			<td style="width:119px;">R0189L</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Phusion polymerase</td>
			<td style="width:145px;">Fisher</td>
			<td style="width:119px;">F-530L</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Qiagen PlasmidPlus Midi kit</td>
			<td style="width:145px;">Qiagen</td>
			<td style="width:119px;">12943</td>
			<td style="width:167px;">contains endotoxin rinse buffer</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">QIAQuick Gel Extraction</td>
			<td style="width:145px;">Qiagen</td>
			<td style="width:119px;">28704&#160;</td>
			<td>for sequencing induced mutations</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Phenol:chloroform:Isoamyl alcohol</td>
			<td style="width:145px;">Sigma</td>
			<td>P2069-100ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sodium acetate</td>
			<td style="width:145px;">Sigma</td>
			<td>S2889-250G</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Ethanol (molecular biology grade)</td>
			<td style="width:145px;">Sigma</td>
			<td>E7023-500ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Agarose</td>
			<td style="width:145px;">Sigma</td>
			<td style="width:119px;">A9539</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">50X Tris-acetate-EDTA buffer</td>
			<td style="width:145px;">ThermoFisher</td>
			<td style="width:119px;">B49</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">0.5-10KbRNA ladder</td>
			<td style="width:145px;">ThermoFisher</td>
			<td style="width:119px;">15623-200</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Nanodrop&#160; Spectrophotometer</td>
			<td style="width:145px;">Thermo Scientific</td>
			<td style="width:119px;">Nanodrop 2000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Paraformaldehyde</td>
			<td style="width:145px;">Sigma</td>
			<td>158127-500G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">10X PBS</td>
			<td style="width:145px;">ThermoFisher</td>
			<td>70011-044</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1kb Plus DNA Ladder</td>
			<td style="width:145px;">ThermoFisher</td>
			<td>10787-018</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Potassium Chloride</td>
			<td style="width:145px;">Sigma</td>
			<td>P9541-500G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Magnesium Chloride</td>
			<td style="width:145px;">Sigma</td>
			<td>M8266-100G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">NP-40</td>
			<td style="width:145px;">ThermoFisher</td>
			<td>28324</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Tween 20</td>
			<td style="width:145px;">Sigma</td>
			<td>P1379-500ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Tris pH 8.3</td>
			<td style="width:145px;">Teknova</td>
			<td>T1083</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">12-strip PCR tube</td>
			<td style="width:145px;">Thermo Scientific</td>
			<td style="width:119px;">AB-1113</td>
			<td></td>
		</tr>
	</tbody>
</table>

microinjection for transgenesis and genome editing in threespine sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

For reporter gene transgenes that have enhancer activity, successful injection will result in specific, cellular expression of the transgene (Figure 4A, 4C). Injected fish can then be outcrossed to produce stable lines (example of a BAC stable line shown in Figure 4B). Injecting DNA into stickleback embryos typically results in far higher lethality than RNA alone. It is typical to see up to 50% (sometimes even more) lethality or malformation (see Figure 4D-F, 4I) after injecting Tol2 reporter constructs (similar to the previously described meganuclease method21). However, the results vary widely based on the specific construct, the embryo morphology, and skill level. For an active enhancer, 40-50% of embryos generally will show tissue-specific transgene expression, for example in the median and pectoral fins (Figure 5). The degree of background and nonspecific fluorescence (Figure 4G-I) varies widely based on the promoter used; the zebrafish hsp70l promoter tends to be leaky, especially in muscle and neural tissue, and BACs tend to have high background expression in the yolk (similar to Figure 4G). The carp beta actin41 promoter is less leaky but also drives considerably fainter GFP expression. Transmission of Tol2 plasmid transgenes can be high, with up to 100% of GFP+ F0 fish producing transgenic offspring (Table 2). However, the percent of offspring carrying the transgene varies widely, from &#60;1% to 72% (Table 2). Saving only GFP+ injected embryos generally increases transmission efficiency. BACs tend to have far lower transgenesis rates, with only up to 10% of F0 injected stickleback embryos showing fluorescence in expected tissues. The transmission rate of BACs is lower than that of plasmid constructs, with only up to 14% of screened stickleback transmitting the BAC (Table 2), which is similar to the reported transmission rate of 15% in zebrafish32.
In contrast to the relatively low efficiency of BAC transgenesis, typically 70-100% of screened fish injected with TALENs have mosaic lesions (in n = 10 injected clutches that were screened for lesions). This number could be lower with a less efficient TALEN pair, and may vary with injection quality. Figure 6 shows a PCR/restriction digestion for 10 embryos from a single clutch injected with TALENs targeting a PvuII cut site within Tfap2a. An uncut amplicon in each of the injected embryos (lanes 1-10) indicates that a portion of the cells in each embryo carry lesions at the target locus, though each embryo is highly mosaic with a significant wild-type cut band. The amplicon from uninjected embryos in lanes 11-12 are fully digested with PvuII. TALEN-induced mutations are readily transmitted to the next generation. With two different TALEN sets, 50% and 90% of screened F0s transmitted lesions to offspring, with 20-90% of offspring carrying lesions in positive clutches (Table 3). While CRISPR/Cas9 efficiency has not been optimized in stickleback, with one CRISPR guide targeting Pitx2, one out of three injected embryos had lesions based on Sanger sequencing of a PCR product of the CRISPR target (the uncut band was sequenced following restriction digestion, Figure 7). The loss of sequence quality at the predicted cut site indicates a mix of molecular lesions are present. Fin clipping adult F0 fish and screening for lesions using a PCR and restriction digestion assay found 10/22 fish with somatic lesions in the fin. A representative subset of these animals are shown in Figure 8; individual #3 has a high percentage of DNA with lesions, while individual #2 has a low percentage of DNA with lesions.
<img alt="Figure 4" src="/files/ftp_upload/54055/54055fig4.jpg" /> 
Figure 4. Examples of injected embryos. (A) Mosaic embryo injected with an enhancer that drives GFP expression in pectoral (asterisk) and median fins (arrowhead) at 5 days post fertilization (dpf). Scale bar = 500 &#181;m. (B) Stable line of a reporter BAC that drives GFP expression in the embryonic heart at 4 dpf. (C) Mosaic embryo injected with a Col2a1a enhancer that drives mCherry expression in the notochord at 4 dpf. The Col2a1a enhancer was cloned from stickleback DNA with primers 5&#39;-CGCTCCTTGAGGGTTTGAGCTG-3&#39; and 5&#39;-ATACTGTGCTCATTTCGGCCGT-3&#39; which amplify the conserved orthologous enhancer reported in Dale and Topczewski 201145. (D) Example of a normally developing injected embryo. (E-F) Examples of malformed embryos with injection trauma; E is lacking the left eye and F has necrotic tissue along the right side (arrowheads). (G) Example of diffuse GFP expression in yolk, likely the result of injection into the yolk rather than the blastomere. (H) Example of non-specific GFP expression in epidermis (arrowhead). (I) Bright, non-specific, granular GFP expression. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54055/54055fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54055/54055fig5.jpg" /> 
Figure 5. Efficiency of reporter construct injection. Ten clutches were injected with a 190 bp stickleback Bmp6 enhancer construct that drives pectoral fin and median fin expression at 5 dpf20. From each clutch, at least 20 embryos were scored for having tissue-specific (pectoral and/or median fin) and/or nonspecific (all other tissues) GFP expression. The percentage of all surviving injected embryos having non-specific and tissue-specific expression is shown as a boxplot. Horizontal lines indicate the first quartile, median and third quartile; whiskers extend to datum within 1.5 IQR (interquartile range) of the first and third quartile. Data are adapted from Erickson et al. 2015. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54055/54055fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/54055/54055fig6.jpg" /> 
Figure 6. PCR and restriction digestion to screen for TALEN-induced lesions. A TALEN pair targeting Tfap2a was generated and injected as described. DNA was prepared as described above from 2 dpf injected embryos and a 297 bp fragment surrounding the TALEN target sequence was PCR amplified by a high fidelity DNA polymerase using primers 5&#39;-GGGTCGTTGACGTGCGAGTAA-3&#39; and 5&#39;-AGCGGGACAACGTCATCACTTA-3&#39;. Lanes 1-10 are injected, lanes 11-12 are uninjected, digested controls, and lanes 13-14 are uninjected, undigested controls. PvuII cuts the wild-type sequence into two approximately equal size bands. Uncut bands indicate presence of molecular lesions in the target sequence. All injected embryos in lanes 1-10 show signs of molecular lesions (undigested bands), however all of the embryos either have monoallelic mutations and/or are mosaic as they also have cut (wild-type) bands. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54055/54055fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/54055/54055fig7.jpg" /> 
Figure 7. Sanger sequencing from mosaic F0 CRISPR/Cas9 injected embryo. A CRISPR guide RNA (5&#39;-GTGGACCAACCTCACGG-3&#39;) against Pitx2 (shown at top) was co-injected with Cas9 mRNA (transcribed from pCS2-nCas9n plasmid as described38) and embryos were screened for lesions using a restriction enzyme assay. The uncut band was gel extracted and sequenced by Sanger sequencing. The sequence quality degrades at the predicted cleavage site (arrow below) due to the mosaic mix of lesions present in the injected embryo. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54055/54055fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" src="/files/ftp_upload/54055/54055fig8.jpg" /> 
Figure 8. Analysis of CRISPR F0 caudal fin clips. DNA was prepared from fin clips from F0 juveniles raised from embryos injected with CRISPRs against Pitx2. The CRISPR/Cas9 target was amplified with primers 5&#39;-CTCGGATGACCCTTCAAAAA-3&#39; and 5&#39;-GGCCCAAATTACCCACATTT-3&#39;, and the product was digested with EcoRI. Four individuals are shown; an uncut PCR product is on the left and digested PCR product is on the right for each numbered individual. Product the size of the uncut band (~230 bp) in the digested lane indicates the presence of a lesion. Individuals 2, 3 and 4 all show signs of a molecular lesion, indicated with asterisks, with varying mosaicism between individuals. Relevant ladder sizes are indicated on left. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54055/54055fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Construct</td>
			<td>GFP offspring/total F0 screened fish (%)</td>
			<td>% F1 offspring positive</td>
			<td>note</td>
		</tr>
		<tr>
			<td>BAC A</td>
			<td>6/46&#160; (13%)</td>
			<td>4-19%</td>
			<td></td>
		</tr>
		<tr>
			<td>BAC B</td>
			<td>1/41 (2%)</td>
			<td>4%</td>
			<td></td>
		</tr>
		<tr>
			<td>BAC C</td>
			<td>5/42 (12%)</td>
			<td>3-40%</td>
			<td></td>
		</tr>
		<tr>
			<td>BAC D</td>
			<td>3/22 (14%)</td>
			<td>5-15%</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid A</td>
			<td>2/38 (5%)</td>
			<td>2-5%</td>
			<td>all F0 embryos screened, not just GFP+</td>
		</tr>
		<tr>
			<td>plasmid B</td>
			<td>3/16 (19%)</td>
			<td>&#60;1-8%</td>
			<td>all F0 embryos screened, not just GFP+</td>
		</tr>
		<tr>
			<td>plasmid C</td>
			<td>1/11 (9%)</td>
			<td>10%</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid D</td>
			<td>2/11 (18%)</td>
			<td>1-45%</td>
			<td>plasmid D injected into 2 genetic backgrounds</td>
		</tr>
		<tr>
			<td>plasmid D</td>
			<td>5/5 (100%)</td>
			<td>16-72%</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid E</td>
			<td>2/3 (67%)</td>
			<td>2-22%</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid F</td>
			<td>2/6 (33%)</td>
			<td>&#60;1-65%</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid G</td>
			<td>3/8 (38%)</td>
			<td>2-56%</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid H</td>
			<td>3/18 (17%)</td>
			<td>not scored</td>
			<td></td>
		</tr>
		<tr>
			<td>plasmid I</td>
			<td>5/24 (21%)</td>
			<td>not scored</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 2: Transgene transmission efficiencies for BACs and enhancer constructs. F0 injected embryos were raised to adulthood and then outcrossed to wild-type fish and the F1 offspring scored for GFP fluorescence. The number of F0 individuals that transmitted the transgene is expressed as a percentage of all screened F0 fish. The range of percentages of F1 offspring carrying the transgene is also shown for those clutches that had GFP positive fish.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>TALEN</td>
			<td>% F0 transmitting lesions</td>
			<td>% F1 offspring positive</td>
		</tr>
		<tr>
			<td>A</td>
			<td>9/10 (90%)</td>
			<td>20-90%</td>
		</tr>
		<tr>
			<td>B</td>
			<td>4/8 (50%)</td>
			<td>20-72%</td>
		</tr>
	</tbody>
</table>
Table 3: Transmission efficiencies for two TALEN pairs. F0 injected embryos were raised to adulthood and then outcrossed to wild-type fish and the F1 offspring screened for TALEN lesions. The percentage of injected individuals transmitting lesions is shown, as well as the range of percentages of offspring with lesions in those clutches that transmitted lesions. TALEN A targeted a Bmp6 enhancer20, TALEN B targeted Tfap2a (unpublished).

Watch this Scientific Journal Video about Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks at JoVE.com

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

microinjection-for-transgenesis-genome-editing-threespine

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

13.8K Views.  University of California, Berkeley. Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

Video: Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

Genome-wide Snapshot of Chromatin Regulators and States in Xenopus Embryos by ChIP-Seq

The recruitment of chromatin regulators and the assignment of chromatin states to specific genomic loci are pivotal to cell fate decisions and tissue and organ formation during development. Determining the locations and levels of such chromatin features in vivo will provide valuable information about the spatio-temporal regulation of genomic elements, and will support aspirations to mimic embryonic tissue development in vitro. The most commonly used method for genome-wide and high-resolution profiling is chromatin immunoprecipitation followed by next-generation sequencing (ChIP-Seq). This protocol outlines how yolk-rich embryos such as those of the frog Xenopus can be processed for ChIP-Seq experiments, and it offers simple command lines for post-sequencing analysis. Because of the high efficiency with which the protocol extracts nuclei from formaldehyde-fixed tissue, the method allows easy upscaling to obtain enough ChIP material for genome-wide profiling. Our protocol has been used successfully to map various DNA-binding proteins such as transcription factors, signaling mediators, components of the transcription machinery, chromatin modifiers and post-translational histone modifications, and for this to be done at various stages of embryogenesis. Lastly, this protocol should be widely applicable to other model and non-model organisms as more and more genome assemblies become available.

Genome-wide Snapshot of Chromatin Regulators and States in Xenopus Embryos by ChIP-Seq

The question of how chromatin regulators and chromatin states affect the genome in vivo is key to our understanding of how early cell fate decisions are made in the developing embryo. ChIP-Seq&#8212;the most popular approach to investigate chromatin features at a global level&#8212;is outlined here for Xenopus embryos.

The recruitment of chromatin regulators and the assignment of chromatin states to specific genomic loci are pivotal to cell fate decisions and tissue and ...

Mosaic Zebrafish Transgenesis for Functional Genomic Analysis of Candidate Cooperative Genes in Tumor Pathogenesis

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers. To robustly and efficiently identify oncogenic &#8220;drivers&#8221; among these tumors and define their complex relationships with concurrent genetic alterations during tumor pathogenesis remains a daunting task. Recently, zebrafish have emerged as an important animal model for studying human diseases, largely because of their ease of maintenance, high fecundity, obvious advantages for in vivo imaging, high conservation of oncogenes and their molecular pathways, susceptibility to tumorigenesis and, most importantly, the availability of transgenic techniques suitable for use in the fish. Transgenic zebrafish models of cancer have been widely used to dissect oncogenic pathways in diverse tumor types. However, developing a stable transgenic fish model is both tedious and time-consuming, and it is even more difficult and more time-consuming to dissect the cooperation of multiple genes in disease pathogenesis using this approach, which requires the generation of multiple transgenic lines with overexpression of the individual genes of interest followed by complicated breeding of these stable transgenic lines. Hence, use of a mosaic transient transgenic approach in zebrafish offers unique advantages for functional genomic analysis in vivo. Briefly, candidate transgenes can be coinjected into one-cell-stage wild-type or transgenic zebrafish embryos and allowed to integrate together into each somatic cell in a mosaic pattern that leads to mixed genotypes in the same primarily injected animal. This permits one to investigate in a faster and less expensive manner whether and how the candidate genes can collaborate with each other to drive tumorigenesis. By transient overexpression of activated ALK in the transgenic fish overexpressing MYCN, we demonstrate here the cooperation of these two oncogenes in the pathogenesis of a pediatric cancer, neuroblastoma that has resisted most forms of contemporary treatment.

The goal of this study is to demonstrate how the mosaic transgenesis strategy can be used in zebrafish to rapidly and efficiently assess the relative contributions of multiple oncogenes in tumor initiation and progression in vivo.

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers.  To robustly and efficiently ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental studies. Like other aphids, A. pisum propagate rapidly via parthenogenetic viviparous reproduction, where the embryos develop within egg chambers in an assembly-line fashion in the ovariole. Previously we have established a robust platform of whole-mount in situ hybridization allowing detection of mRNA expression in the aphid embryos. For analyzing the expression of protein, though, established protocols for immunostaining the ovarioles of asexual viviparous aphids did not produce satisfactory results. Here we report conditions optimized for increasing tissue permeability and decreasing background staining, both of which were problems when applying established approaches. Optimizations include: (1) incubation of proteinase K (1 &#181;g/ml, 10 min), which was found essential for antibody penetration in mid- and late-stage aphid embryos; (2) replacement of normal goat serum/bovine serum albumin with a blocking reagent supplied by a Digoxigenin (DIG)-based buffer set and (3) application of methanol rather hydrogen peroxide (H2O2) for bleaching endogenous peroxidase; which significantly reduced the background staining in the aphid tissues. These critical conditions optimized for immunostaining will allow effective detection of gene products in the embryos of A. pisum and other aphids.

A protocol of whole-mount immunostaining on aphid embryos is presented, in which critical conditions for decreasing background staining and increasing tissue permeability are addressed. These conditions are specifically developed for effective detection of protein expression in the embryonic tissues of aphids.

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of cells or tissues in model organisms. These systems contain two components as separate transgenic lines. A driver line expresses a transcriptional activator under the control of tissue-specific promoters/enhancers, and a reporter/effector line harbors a target gene placed downstream to the binding site of the transcription activator. Animals harboring both components induce tissue-specific transactivation of a target gene expression. Precise spatiotemporal expression of the gene in targeted tissues is critical for unbiased interpretation of cell/gene activity. Therefore, developing a method for generating exclusive cell/tissue-specific driver lines is essential. Here we present a method to generate highly tissue-specific targeted expression system by employing a &#34;Clustered Regularly Interspaced Short Palindromic Repeat/CRISPR-associated&#34; (CRISPR/Cas)-based genome editing technique. In this method, the endonuclease Cas9 is targeted by two chimeric guide RNAs (gRNA) to specific sites in the first coding exon of a gene in the Drosophila genome to create double-strand breaks (DSB). Subsequently, using an exogenous donor plasmid containing the transactivator sequence, the cell-autonomous repair machinery enables homology-directed repair (HDR) of the DSB, resulting in precise deletion and replacement of the exon with the transactivator sequence. The knocked-in transactivator is expressed exclusively in cells where the cis-regulatory elements of the replaced gene are functional. The detailed step-by-step protocol presented here for generating a binary transcriptional driver expressed in Drosophila fgf/branchless-producing epithelial/neuronal cells can be adopted for any gene- or tissue-specific expression.

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Here, we present a method for generating tissue-specific binary transcription systems in Drosophila by replacing the first coding exon of genes with transcription drivers. The CRISPR/Cas9-based method places a transactivator sequence under the endogenous regulation of a replaced gene, and consequently facilitates transctivator expression exclusively in gene-specific spatiotemporal patterns.

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...

Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica

The firebrat Thermobia domestica is an ametabolous, wingless species that is suitable for studying the developmental mechanisms of insects that led to their successful evolutionary radiation on the earth. The application of genetic tools such as genome editing is the key to understanding genetic changes that are responsible for evolutionary transitions in an Evo-Devo approach. In this article, we describe our current protocol for generating and maintaining mutant strains of T. domestica. We report a dry injection method, as an alternative to the reported wet injection method, that allows us to obtain stably high survival rates in injected embryos. We also report an optimized environment setting to mate adults and obtain subsequent generations with high efficiency. Our method underlines the importance of taking each species&#8217; unique biology into account for the successful application of genome editing methods to non-traditional model organisms. We predict that these genome editing protocols will help in implementing T. domestica as a laboratory model and to further accelerate the development and application of useful genetic tools in this species.

Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica

We provide a detailed protocol for rearing, microinjection of eggs and for efficient mating of the firebrat Thermobia domestica to generate and maintain mutant strains after genome editing.

The firebrat Thermobia domestica is an ametabolous, wingless species that is suitable for studying the developmental mechanisms of insects that led to ...

Brain Organoid Generation from Induced Pluripotent Stem Cells in Home-Made Mini Bioreactors

The iPSC-derived brain organoid is a promising technology for in vitro modeling the pathologies of the nervous system and drug screening. This technology has emerged recently. It is still in its infancy and has some limitations unsolved yet. The current protocols do not allow obtaining organoids to be consistent enough for drug discovery and preclinical studies. The maturation of organoids can take up to a year, pushing the researchers to launch multiple differentiation processes simultaneously. It imposes additional costs for the laboratory in terms of space and equipment. In addition, brain organoids often have a necrotic zone in the center, which suffers from nutrient and oxygen deficiency. Hence, most current protocols use a circulating system for culture medium to improve nutrition.
Meanwhile, there are no inexpensive dynamic systems or bioreactors for organoid cultivation. This paper describes a protocol for producing brain organoids in compact and inexpensive home-made mini bioreactors. This protocol allows obtaining high quality organoids in large quantities.

Here we describe a protocol for generating brain organoids from human induced pluripotent stem cells (iPSCs). To obtain brain organoids in large quantities and of high quality, we use home-made mini bioreactors.

The iPSC-derived brain organoid is a promising technology for in vitro modeling the pathologies of the nervous system and drug screening. This technology ...