Name: generation of transgenic hydra by embryo microinjection
Uploaded: 2014-09-11T15:00:00.000+00:00
Duration: 9 min 10 s
Description: Watch this Scientific Journal Video about Generation of Transgenic Hydra by Embryo Microinjection at JoVE.com

This work was supported by a G. Harold &#38; Leila Y. Mathers Award to H.L. and an NIH grant (R24 GM080527) to R.E.S. C.E.J. was an NRSA Postdoctoral Fellow (NIH F32GM9037222) and is currently supported by a Mentored Research Scientist Development Award from the National Institute on Aging (K01AG04435). We would like to thank the reviewers for helpful comments.

1. Preparation of Plasmid DNA, Needles, and Injection Dishes
<ol>
	<li>Prepare the plasmid DNA using a High Speed Maxi or Midi kit and concentrate the DNA to 2.5 mg/ml using ethanol precipitation. The DNA pellet should be dissolved in nuclease-free deionized water. Store the DNA in 10-20 &#181;l aliquots at -20&#176;C. (See Supplementary Table 1 for a list of available vectors)</li>
	<li>Make an injection dish using agarose to construct a trough for holding embryos in a 100 x 15 mm Petri dish.
	<ol>
		<li>Place a 75 x 50 glass microscope slide into a 100 x 15 mm Petri dish at an angle. Pour approximately 50 ml of 2% agarose melted in Hydra medium24 into the Petri dish. Alternatively, pour the agarose into the dish first and float a &#8220;Microinjection Fish Mold&#8221; on top.</li>
		<li>After the agarose has solidified, remove the glass slide or the mold. If a glass slide was used, cut away excess agarose with a razor blade to form a wall. Pour Hydra medium into the dish and use it immediately or store it at 4&#176;C until needed. Note: Injection dishes can be reused if stored at 4&#176;C between uses.</li>
	</ol>
	</li>
	<li>Pull needles from borosilicate glass capillaries with a filament on the micropipette puller using the following conditions: heat 525, pull 75, velocity 100, time 50. Store the needles by pressing them into a narrow strip of clay in a Petri dish such that they are held parallel to the bottom of the dish.</li>
	<li>To prepare the injection solution, combine 3 &#181;l of 2.5 mg/ml plasmid DNA solution with 2 &#181;l of 10% phenol red. Vortex the solution briefly and then spin it for 10 min&#160;at maximum speed in a microcentrifuge to remove any particles that might clog the needle.</li>
	<li>Under a dissecting microscope, clip the needle a few millimeters from the tip with forceps to create a small opening. Note: There is some flexibility in the appropriate size of the opening because the pressure can be adjusted in step 3.3 to accommodate this variation.</li>
	<li>Place the Petri dish so that it is holding the needle in a vertical position with the tip down. Load the needle with approximately 0.5 &#181;l of injection solution by pipetting the liquid into the back end of the needle. Allow capillary action to pull the solution into the tip of the needle.</li>
</ol>
2. Preparation of Embryos for Injection
<ol>
	<li>On a daily basis scan the Hydra AEP strain culture for polyps producing eggs. Collect these polyps and set them aside in a separate culture dish. Observe these polyps every few hours during the day to monitor the progress of egg formation. When eggs break through the ectoderm and sit on a ring of retracted ectodermal cells they are ready for injection25. Place these polyps in a dish with several Hydra that have testes for at least 1 hr before injection to allow for fertilization. Hydra males will release sperm without any special manipulation.</li>
	<li>If females are found in the AEP colony with fully-formed eggs, which are approximately 400 &#956;m in diameter25, or 1- to 8-cell stage embryos, also collect these for injection. Immediately inject embryos that have started cleaving. Note: It is not possible to tell the difference between fully-formed eggs and single-celled zygotes, thus these should be incubated with males prior to injection.</li>
	<li>Before injection, remove most of the parental tissue above and below the embryo using a scalpel with a #15 blade. Leave only the embryo with a small piece of body column attached, to be used to manipulate the embryo with forceps if necessary. Note: The tissue that is dissected away from the embryo will regenerate and should be saved to ensure that females remain in the Hydra colony.</li>
	<li>Using a Pasteur pipette, move embryos to the injection dish, arranging them parallel to the wall of the agarose trough.</li>
</ol>
3. Microinjection of Plasmid DNA
<ol>
	<li>Mount the microinjector on a magnetic stand that sits on an iron plate. Mount the injection holder, which is the part of the microinjector that holds the needle, on a joystick micromanipulator. Mount the joystick manipulator to the iron plate, with a magnetic stand. Place this entire set-up on the right side of a dissecting microscope and position the injection holder so that it is visible in the observation field.</li>
	<li>Place the injection dish with embryos under the dissecting microscope, oriented such that the vertical wall of the agarose trough is to the left. Insert the needle into the injection holder and lower the tip of the needle into the Hydra medium in the dish.</li>
	<li>Slowly turn the knob of the syringe in the clockwise direction until mineral oil fills the top of the needle and a steady stream of injection solution is observed exiting the needle into the Hydra medium. If the stream is too strong, lower the pressure by turning the knob counterclockwise. Practice this step to routinely obtain a suitable pressure, which depends on the size of the needle opening (i.e., how the needle was clipped in step 1.5). Note: If the stream is too strong, the embryo will not survive the injection.</li>
	<li>While viewing through the dissecting microscope, move the injection dish so that the first embryo is at the center of the field. Move the needle so that it is touching that embryo. Use the micromanipulator to pierce the embryo with the needle. Note: The vertical wall of the agarose trough will keep the embryo from being pushed aside by the needle.</li>
	<li>Allow the injection solution to flow into the embryo 1 or 2 sec and then remove the needle quickly. If the embryo has more than one cell, inject each cell individually. Use forceps to reorient the embryo to align the next cell with the tip of the needle.</li>
	<li>After the first embryo is injected, move the dish so that the next embryo is in the injection position and repeat the procedure; continue until all embryos have been injected.</li>
</ol>
4. Culturing and Hatching of Embryos
<ol>
	<li>Move all of the injected embryos into a dish with a few Hydra that have testes (this is to ensure that all embryos are fertilized). Incubate the embryos in an 18 &#176;C incubator while they continue embryogenesis.</li>
	<li>Once the cuticle stage is reached, move each injected embryo to an individual well of a 24-well plate filled with Hydra medium.</li>
	<li>Incubate the injected embryos for 2 weeks in the dark at 18 &#176;C.</li>
	<li>After the 2 week incubation period, check each injected embryo under the dissecting microscope. Note any cases in which the cuticle-stage embryo has detached from the small piece of parental tissue and the parental tissue has regenerated. Remove this regenerated polyp immediately so that it is not mistaken for a new hatchling.</li>
	<li>Move the dish of injected embryos under an aquarium light at RT. Check the plates each day and collect hatchlings. New hatchlings will be white because the typical pink color of Hydra polyps comes from the Artemia they eat.</li>
	<li>Observe the hatchlings under a fluorescence microscope to detect transgene expression.</li>
</ol>

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The authors have nothing to disclose

Hydra routinely reproduces asexually, but requires environmental stimuli to begin producing gametes. These stimuli are not well-defined for most Hydra species and may differ from strain to strain. A significant hurdle to the production of transgenic Hydra is obtaining embryos on a regular basis because it can be difficult in a laboratory setting to induce Hydra to become sexual. The AEP strain25, however, produces gametes readily in the laboratory and this is the only strain...

Hydra has been used to study regeneration, pattern formation, and stem cells for approximately 250 years2. Hydra has a simple body plan consisting of three cell lineages: ectodermal epithelial, endodermal epithelial, and interstitial. The tubular body is formed by the ectodermal and endodermal epithelial lineages, each of which is a single cell layer. All of the epithelial cells in the body column are mitotic. When epithelial cells are displaced into the extremities3, the head (mouth and tentacles) at the oral end or the foot (basal disc) at the aboral end, they arrest in the G2 phase of the cell cycle and change cell fate4. The cells of the interstitial lineage reside within the interstices between the epithelial cells. This lineage is supported by multipotent stem cells that are located in the ectodermal epithelial layer of the body column5. The interstitial stem cells give rise to three somatic cell types (nerves, gland cells, and nematocytes) and the germ cells6,7.
As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among multicellular animals. Until recently, these efforts were impeded by the lack of reliable methods for the perturbation of gene function. However, with the development of transgenic methodology1, we are now able to take full advantage of Hydra to gain a better understanding of the basic mechanisms common to multicellular animals, such as stem cell function, regeneration, and patterning. Transgenic Hydra lines are established by injection of plasmid DNA into embryos, which results in random integration and chimeric expression in a substantial frequency of hatchlings. A line with uniform expression in a particular lineage can be established by asexual propagation. The ability to clonally propagate transgenic Hydra lines is an advantage over the majority of animal models, which can be propagated only by sexual reproduction. In addition, transgenic cells can be tracked easily in vivo due to the transparency of the animal and the absence of endogenous fluorescent proteins 8.
In the seven years since the first transgenic Hydra lines were made1, such lines have been used for a variety of applications. Expression of fluorescent proteins in different cell types has made it possible to track cell movement, observe changes in cell shape, and track cell fates both in wild type conditions and after chemical perturbation1,5,9-12. In addition, expression of different fluorescent proteins in the various lineages allows for FACS isolation of specific cell populations. This technique has been used for the sequencing of stem-cell specific mRNAs and lineage-specific small RNAs13,14. While the promoter of one of the two Hydra actin genes has been most widely used, a few cell-type specific promoters have been identified and used to drive expression of GFP in transgenic Hydra9,11,15,16. In the future, cell-type specific promoters will allow for the observation and collection of any specific cell type. In addition, a transgenic approach was successfully used to define the cis-acting regulatory elements of the Wnt3 promoter17.
The development of transgenic methods in Hydra provides a robust approach for testing the function of genes by ectopic expression, overexpression, and knockdown. Transgenic animals have been made that express fluorescently-tagged proteins in order to examine both function and cellular localization18-20. In addition, the expression of RNA hairpins in the 3&#8217;UTR of a GFP transgene leads to knockdown of target genes21,22. In these approaches GFP is required to identify and track the transgenic tissue during the creation of the transgenic line. However, it is likely that in some cases the GFP molecule would interfere with the function of the tagged protein. A recent study demonstrates that Hydra genes can be arranged in an operon configuration, i.e.,&#160;polycistronic transcripts are made, which are then separated by trans-spliced leader addition and translated separately23. By placing a gene encoding a protein or an RNA hairpin in the upstream position of an operon and a fluorescent protein gene in the downstream position, one can track transgenic tissue without having to tag the gene encoding the protein or RNA hairpin. This method has been used to express an RNA hairpin in an operon configuration with DsRed2 in order to achieve gene knockdown14.

<table><tbody><tr><td>AEP Hydra Strain</td><td></td><td></td><td>Email: Celina Juliano at celina.juliano@yale.edu or Hiroshi Shimizu bubuchin2006@yahoo.co.jp</td></tr><tr><td>High Speed Maxi Kit</td><td>Qiagen</td><td>12662</td><td></td></tr><tr><td>100 x 15 mm Petri Dishes</td><td>BD Falcon</td><td>351029</td><td></td></tr><tr><td>75 x 50 mm Glass Microscope Slide</td><td>Sigma</td><td>CLS294775X50</td><td></td></tr><tr><td>Microinjection Fish Mold</td><td>IBI Scientific</td><td>FM-600</td><td></td></tr><tr><td>Borosilicate Glass with Filament</td><td>Sutter Instrument</td><td>BF100-50-10</td><td>O.D.: 1.0 mm, I.D.: 0.50 mm, 10 cm length</td></tr><tr><td>Flaming/Brown Micropipette Puller</td><td>Sutter Instrument</td><td>P-97</td><td></td></tr><tr><td>Phenol Red</td><td>Sigma</td><td>P3532</td><td></td></tr><tr><td>Jewelers Forceps, Durmont #5</td><td>Sigma</td><td>F6521</td><td></td></tr><tr><td>Scalpel Blade #15</td><td>Fisher</td><td>50-822-421</td><td></td></tr><tr><td>Mineral oil</td><td>Sigma</td><td>M8410-500ML</td><td></td></tr><tr><td>Microinjector</td><td>Narishige</td><td>IM-9B</td><td></td></tr><tr><td>Magnetic Stand</td><td>Narishige</td><td>GJ-1</td><td></td></tr><tr><td>Iron Plate</td><td>Narishige</td><td>IP</td><td></td></tr><tr><td>Joystick Micromanipulator</td><td>Narishige</td><td>MN-151</td><td></td></tr></tbody></table>

generation of transgenic hydra by embryo microinjection

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among multicellular animals. Hydra is used as a model for the study of regeneration, pattern formation, and stem cells. However, research efforts have been hampered by lack of a reliable method for gene perturbations to study molecular function. The development of transgenic methods has revitalized the study of Hydra biology1. Transgenic Hydra allow for the tracking of live cells, sorting to yield pure cell populations for biochemical analysis, manipulation of gene function by knockdown and over-expression, and analysis of promoter function. Plasmid DNA injected into early stage embryos randomly integrates into the genome early in development. This results in hatchlings that express transgenes in patches of tissue in one or more of the three lineages (ectodermal epithelial, endodermal epithelial, or interstitial). The success rate of obtaining a hatchling with transgenic tissue is between 10% and 20%. Asexual propagation of the transgenic hatchling is used to establish a uniformly transgenic line in a particular lineage. Generating transgenic Hydra is surprisingly simple and robust, and here we describe a protocol that can be easily implemented at low cost.

Establishing transgenic Hydra lines
Feed transgenic hatchlings every 2-3 days with Artemia nauplii. Hatchlings sometimes do not eat for a day or two after hatching. Some new hatchlings will never eat, and thus will not survive. If the hatchling is transgenic in either the ectodermal (Figure 1A) or endodermal epithelial tissue, allow the animal to bud and collect the buds that have the most transgenic tissue (Figure 1B, C). Continue to d...

Watch this Scientific Journal Video about Generation of Transgenic Hydra by Embryo Microinjection at JoVE.com

Generation of Transgenic Hydra by Embryo Microinjection

Stably transgenic Hydra are made by microinjection of plasmid DNA into embryos followed by random genomic integration and asexual propagation to establish a uniform line. Transgenic Hydra are used to track cell movements, overexpress genes, study promoter function, or knock down gene expression using RNAi.

Generation of Transgenic Hydra by Embryo Microinjection

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among ...

generation-of-transgenic-hydra-by-embryo-microinjection

Research

JoVE Journal

Biology

14.0K Views.  Yale University School of Medicine. Stably transgenic Hydra are made by microinjection of plasmid DNA into embryos followed by random genomic integration and asexual propagation to establish a uniform line. Transgenic Hydra are used to track cell movements, overexpress genes, study promoter function, or knock down gene expression using RNAi. 

Video: Generation of Transgenic Hydra by Embryo Microinjection

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

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

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

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

Developmental Biology

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex determination, venom synthesis, and host-symbiont interactions, among others. A major limitation of working with this organism is the lack of effective protocols to perform directed genome modifications. An important part of genome modification is delivery of editing reagents, including CRISPR/Cas9 molecules, into embryos through microinjection. While microinjection is well established in many model organisms, this technique is particularly challenging to perform in N. vitripennis primarily due to its small embryo size, and the fact that embryonic development occurs entirely within a parasitized blowfly pupa. The following procedure overcomes these significant challenges while demonstrating a streamlined, visual procedure for effectively removing wasp embryos from parasitized host pupae, microinjecting them, and carefully transplanting them back into the host for continuation and completion of development. This protocol will strongly enhance the capability of research groups to perform advanced genome modifications in this organism.

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Microinjection of Nasonia vitripennis embryos is an essential method for generating heritable genome modifications. Described here is a detailed procedure for microinjection and transplantation of Nasonia vitripennis embryos, which will greatly facilitate future genome manipulation in this organism.

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex ...

Genetics

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how genes regulate social traits, we have developed mutant ants via delivery of CRISPR complex into young embryos during their syncytial stage. Here, we provide a protocol of CRISPR-mediated mutagenesis in Harpegnathos saltator, a ponerine ant species that displays striking phenotypic plasticity. H. saltator ants are readily reared in a laboratory setting. Embryos are collected for microinjection with Cas9 proteins and in vitro synthesized small guide RNAs (sgRNAs) using home-made quartz needles. Post-injection embryos are reared outside the colony. Following emergence of the first larva, all embryos and larvae are transported to a nest box with a few nursing workers for further development. This protocol is suitable for inducing mutagenesis for analysis of caste-specific physiology and social behavior in ants, but may also be applied to a broader spectrum of hymenopterans and other insects.

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

Many characteristics of insect eusociality rely on within-colony communication and division of labor. Genetic manipulation of key regulatory genes in ant embryos via microinjection and CRISPR-mediated mutagenesis provides insights into the nature of altruistic behavior in eusocial insects.

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how ...

phiC31 Integrase-Mediated Embryo Transgenesis in Drosophila

Embryo Microinjection for Transgenesis in Drosophila

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the construction of transgenic flies. Microinjection requires some types of equipment, including a microinjector, a micromanipulator, an inverted microscope, and a stereo microscope.&#160;Plasmids isolated with a plasmid miniprep kit are qualified for microinjection. Embryos at the pre-blastoderm or syncytial blastoderm stage, where nuclei share a common cytoplasm, are subjected to microinjection. A cell strainer eases the process of dechorionating embryos. The optimal time for dechorionation and desiccation of embryos needs to be determined experimentally. To increase the efficiency of embryo microinjection, needles prepared by a puller need to be beveled by a needle grinder. In the process of grinding needles, we utilize a foot air pump with a pressure gauge to avoid the capillary effect of the needle tip. We routinely inject 120-140 embryos for each plasmid and obtain at least one transgenic line for around 85% of plasmids. This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and&#160;presents a detailed protocol for embryo microinjection for transgenesis in Drosophila.

Author Spotlight: Optimizing Embryo Microinjection for Transgenesis in Drosophila

This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and presents an optimized protocol for embryo microinjection,&#160;a crucial step for creating transgenic flies.

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the ...

Generation and Long-term Maintenance of Nerve-free Hydra

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The interstitial cells can be eliminated through two consecutive treatments with colchicine, a plant-derived toxin that kills dividing cells, thus erasing the potential for renewal of the differentiated cells that are derived from the interstitial stem cells. This allows for the generation of Hydra that lack nerve cells. A nerve-free polyp cannot open its mouth to feed, egest, or regulate osmotic pressure. Such animals, however, can survive and be cultured indefinitely in the laboratory if regularly force-fed and burped. The lack of nerve cells allows for studies of the role of the nervous system in regulating animal behavior and regeneration. Previously published protocols for nerve-free Hydra maintenance involve outdated techniques such as mouth-pipetting with hand-pulled micropipette tips to feed and clean the Hydra. Here, an improved protocol for maintenance of nerve-free Hydra is introduced. Fine-tipped forceps are used to force open the mouth and insert freshly killed Artemia. Following force-feeding, the body cavity of the animal is flushed with fresh medium using a syringe and hypodermic needle to remove undigested material, referred to here as "burping". This new method of force-feeding and burping nerve-free Hydra through the use of forceps and syringes eliminates the need for mouth-pipetting using hand-pulled micropipette tips. It thus makes the process safer and significantly more time efficient. To ensure that the nerve cells in the hypostome have been eliminated, immunohistochemistry using anti-tyrosine-tubulin is conducted.

Generation and Long-term Maintenance of Nerve-free Hydra

Through a double treatment with colchicine, a plant-derived toxin that kills dividing cells, nerve-free Hydra vulgaris can be generated. These Hydra cannot feed or egest on their own. This paper describes an improved method for long-term maintenance of nerve free Hydra vulgaris in the laboratory.

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The ...

Neuroscience

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

The genus Strongyloides consists of multiple species of skin-penetrating nematodes with different host ranges, including Strongyloides stercoralis and Strongyloides ratti. S.&#160;stercoralis is a human-parasitic, skin-penetrating nematode that infects approximately 610 million people, while the rat parasite S.&#160;ratti is closely related to S.&#160;stercoralis and is often used as a laboratory model for S.&#160;stercoralis. Both S.&#160;stercoralis and S.&#160;ratti are easily amenable to the generation of transgenics and knockouts through the exogenous nucleic acid delivery technique of intragonadal microinjection, and as such, have emerged as model systems for other parasitic helminths that are not yet amenable to this technique.
Parasitic Strongyloides adults inhabit the small intestine of their host and release progeny into the environment via the feces. Once in the environment, the larvae develop into free-living adults, which live in feces and produce progeny that must find and invade a new host. This environmental generation is unique to the Strongyloides species and similar enough in morphology to the model free-living nematode Caenorhabditis elegans that techniques developed for C. elegans can be adapted for use with these parasitic nematodes, including intragonadal microinjection. Using intragonadal microinjection, a wide variety of transgenes can be introduced into Strongyloides. CRISPR/Cas9 components can also be microinjected to create mutant Strongyloides larvae. Here, the technique of intragonadal microinjection into Strongyloides, including the preparation of free-living adults, the injection procedure, and the selection of transgenic progeny, is described. Images of transgenic Strongyloides larvae created using CRISPR/Cas9 mutagenesis are included. The aim of this paper is to enable other researchers to use microinjection to create transgenic and mutant Strongyloides.

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

The functional genomic toolkit for the parasitic nematodes Strongyloides stercoralis and Strongyloides ratti includes transgenesis, CRISPR/Cas9-mediated mutagenesis, and RNAi. This protocol will demonstrate how to use intragonadal microinjection to introduce transgenes and CRISPR components into S.&#160;stercoralis and S.&#160;ratti.

The genus Strongyloides consists of multiple species of skin-penetrating nematodes with different host ranges, including Strongyloides stercoralis and ...

Generation of Transgenic Hydra by Embryo Microinjection

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

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

Embryo Microinjection for Transgenesis in Drosophila

Generation and Long-term Maintenance of Nerve-free Hydra

High Throughput Microinjections of Sea Urchin Zygotes

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection