This work was supported by the National Science Foundation IOS 0844948 and IOS 1024811

Keep all sea water or artificial seawater (SW), adult animals, and cultures at 15 &#176;C as much as is practical. Ensure eggs and zygotes are kept immersed in SW.
Commercially prepared sea salts reconstituted with distilled or reverse osmosis water serves well as a source of SW. Check salinity using a hydrometer and adjust salts or water to achieve optimum levels. Keep specific gravity levels between 1.020 to 1.025. Keep all glassware and plasticware separate from all other labware to avoid any contamination with chemicals. Clean embryo grade labware by rinsing with deionized water or occasional soaking in dilute sodium hypochlorite followed by rinsing several times in water.
1. Obtaining and Maturing Gametes from Patiria miniata Adults
<ol>
	<li>Select an animal to excise gonads. Determine sex after excision by looking for the presence of ovaries or testes because P. miniata are not sexually dimorphic.</li>
	<li>Cut a small opening, no larger than 1 cm, along the side of a sea star arm with a scalpel blade (Figure 1A). Using small blunt-ended forceps pull back edges of the incision to access the gonad tissue and pull gonad tissue out through the incision. 
	NOTE: Avoid pulling out gut tissue, which is a loose gray or brown tissue (Figure 1B). Ovaries are typically orange, yellow, or light brown (Figure 1C), while testes are white or beige and when disrupted will cause sea water to become milky or cloudy in appearance (Figure 1D).</li>
	<li>Return females to the aquaria. Isolate males for an hour or two in a container of SW since the stress from handling may induce spawning. Animals heal along the incision site and continue to provide gametes for several months.</li>
	<li>Collect testes into a 1.5 ml Eppendorf tube with as little SW as possible and store on ice until use. Store any testes not used on the day of collection at 4 &#176;C for up to a week.</li>
	<li>Collect ovaries in a small glass culture dish in a several milliliters of SW. To release the oocytes from the ovary tissue, tease apart the tissue with two pairs of forceps until no large pieces remain. 
	NOTE: Optimally, the majority of the oocytes are fully developed. Such oocytes are large, round, and clear yellow or light brown with a distinct germinal vesicle in the oocyte (Figure 2A), compared to the subset of undeveloped oocytes that are smaller, brown, grainy, and opaque (Figure 2B). If greater than around 25% of the oocytes are of the undeveloped variety, select a different female to obtain oocytes.</li>
	<li>Pour the oocytes and remaining tissue through a mesh filter cup with a 200 &#956;m pore size and collect flow through. This will contain all varieties of oocytes but remove ovarian tissue. Dislodge remaining oocytes from the tissue caught by the filter by spraying with SW from a squirt bottle. Filter cups are made using a 50 ml conical tube (Figure 3A-A&#8217;).</li>
	<li>Pour the flow through from step 1.6 through a 100 &#956;m mesh filter cup. Discard small oocytes in flow through. Large fully developed oocytes that are ready for maturation will stay on the filter. Rinse the oocytes into a clean 200 ml beaker.</li>
	<li>Add 95 ml of SW to the 200 ml glass beaker of oocytes. Add 5 ml of 200 &#956;M 1-methyladenine for a final concentration of 10 &#956;M.</li>
	<li>Transfer the oocytes to a large shallow culture dish and place at 15 &#176;C. A high surface area to volume culture dish allows for oxygen exchange. The oocytes do not mature or may not develop well if they are overcrowded during this maturation phase. Split into several dishes of SW plus 10 &#956;M 1-methlyadenine until culture is less than 200 oocytes/ml.</li>
	<li>Allow the oocytes to mature for 45-90 min, but no longer as this will render the oocytes unable to be fertilized. To determine if maturation is complete, look at the oocytes under a dissecting scope. The germinal vesicle moves towards and fuses with the cell membrane during maturation. It then breaks down and therefore is no longer visible in mature oocytes (Figure 2C).</li>
</ol>
2. Fertilization of Mature Oocytes
<ol>
	<li>Chose a small cluster of testis tissue, roughly pea-sized, and mince in a small glass culture dish with approximately 1 ml of SW.</li>
	<li>Use a Pasteur pipette to transfer 2-3&#160;drops of the resulting milky sea water to around 100 ml of the mature oocytes and swirl the culture dish to mix. Avoid transferring pieces of testis tissue. Avoid adding greater quantities of sperm as this may result in polyspermy and subsequent poor development.</li>
	<li>After several minutes observe the oocytes under a dissecting scope. Fertilized eggs, or zygotes, have a fertilization envelope, which is a clear bubble that rises off the cell membrane (Figure 2D). 
	NOTE: Do not proceed with the culture if less than around 90% of eggs fertilize since poor fertilization rates are an indication of an unhealthy culture that may also not develop well.</li>
	<li>After fertilization is complete, change the SW on the zygotes by pouring through a 100 &#956;m mesh filter and rinsing into a culture dish with fresh SW. It is important to remove the excess sperm to prevent oxygen starvation. Place the culture dish at 15 &#176;C for 15-30 min.</li>
</ol>
3. De-jellying and Rowing Fertilized Eggs
<ol>
	<li>Prepare injection dishes by taking the lid from a 60 x 15 mm polystyrene Petri dish.
	<ol>
		<li>Draw a permanent marker pen line on the back of the dish, around the area that matches the field of view on the microinjection microscope. This ensures that all rowed embryos are within the microscope field of view.</li>
		<li>Use a glass Pasteur pipette to score a line through the circle, preferably off to one side of the circle (Figure 3B). Use this line later to break injection needles (Step 4.5). 
		NOTE: Reagents commonly used to adhere sea urchin embryos, e.g., protamine sulfate or poly-L Lysine, are unnecessary. De-jellied P. miniata zygotes stick very well to uncoated plastic. Note that they do not stick well to glass dishes.</li>
	</ol>
	</li>
	<li>Add around 10 ml SW to each dish so that the surface is covered, but the water will not splash excessively in the dish as this may perturb rowed embryos.</li>
	<li>Remove the P. miniata zygote jelly coat with acid SW prior to rowing. This jelly coat is more extensive than that observed for model sea urchin species.
	<ol>
		<li>Prepare acid SW in the range of pH 4.0-4.2. The pH is critical to allow efficient removal of jelly coat while permitting normal development.</li>
		<li>Add single drops of 1 N (1 M) HCl in SW to a 1 L stock of SW. Allow each drop to mix completely and monitor pH before adding more HCl. 
		NOTE: One to several ml of HCl results in the appropriate pH, but do not add the entire volume at once. 
		NOTE: Prepare 1 N HCl in fume hood with SW and 12 N (12 M) HCl.</li>
		<li>Remove SW on zygotes by pouring them onto a 100 &#956;m mesh filter. Rinse into a 200 ml beaker in the smallest amount of SW possible (approximately 10 ml). Fill the beaker up to 150 ml with acid SW (pH 4.0) and allow the zygotes to sit in the acid SW for 3 min.</li>
	</ol>
	</li>
	<li>Strip jelly coat from the zygotes by pouring them back and forth between two 200 ml beakers, each time passing the zygotes through a 200 &#956;m mesh filter. Pour the zygotes through the filter around 5-10x over about a 2 min time window.</li>
	<li>Remove acid SW by pouring the zygotes onto a 100 &#956;m mesh filter. Rinse the zygotes into a small glass culture dish with SW. De-jellied zygotes may stick to plastic dishes. Row embryos immediately; they start to produce more jelly coat which leads to poor adhesion to plastic dishes over time.</li>
	<li>Use a mouth pipette (Figure 3C-C&#8217;) to pick up zygotes from the glass culture dish and place them into straight lines on the injection dishes. Melt the center of the thin side of a Pasteur pipette over a flame until the glass is soft. Quickly pull the ends of the glass apart to produce a very thin stretch of glass. Once the glass is cool, break off the small end of the Pasteur pipette (for additional information on preparing a mouth pipette see Wessel et al. 200417 or Sweet et al. 200418).
	<ol>
		<li>Make a rowing pipette such that the diameter is just larger than the diameter of the embryos so that only a single zygote will enter and exit the pipette at a time. This will allow a single row to form. Smaller diameter pipettes may strip the fertilization envelope and disrupt the zygotes as P. miniata zygotes do not have a hyaline membrane and are quite soft.</li>
		<li>If the zygotes do not stick to the plastic dish, keep them for additional time in acid SW (up to several more minutes). Alternatively a smaller diameter pipette may aid in removing jelly coat to allow the zygotes to stick more effectively.</li>
		<li>The zygotes are slightly positively buoyant and tend to float or hover just above the bottom of the dish. In this case, position the rowing pipette so that the zygotes are gently pressed onto the surface of the dish as they exit the pipette (Figure 3D). 
		NOTE: It is permissible to fit many rows on these dishes. P. miniata embryonic membranes do not become more difficult to penetrate over time and therefore one can inject hundreds of zygotes on a single dish.</li>
	</ol>
	</li>
</ol>
4. Injection of Zygotes
<ol>
	<li>Prepare injection solutions with appropriate concentrations of morpholino antisense oligonucleotides (MASO), mRNA, or DNAs in a final concentration of 200 mM KCl. 400-800 &#956;M MASO and 2.5 ng/&#956;l of DNA tend to produce morphant phenotypes while minimizing toxicity. The final concentration in the egg will be 1/125th the starting concentration14. For injection solutions containing MASOs, heat to 65 &#176;C for 5 min immediately prior to use and then store at room temperature.
	<ol>
		<li>For ease of sorting injected embryos, add rhodamine dextran tracer to the injection solution to obtain a concentration of 0.1%.</li>
	</ol>
	</li>
	<li>Prepare injection needles by pulling glass capillary tubing (1.0 mm OD x 0.75 mm ID, 100 mm length) in a needle pulling instrument following manufacturer&#8217;s instructions. Needles suitable for injection in sea urchins also work well for sea stars, so use similar parameters19.</li>
	<li>Load approximately 2 &#956;l of injection solution into an injection needle using a microloader pipette tip.</li>
	<li>Insert the blunt end of the needle into the manipulator. Set picospritzer to 100 msec pulse and an air pressure of 40 PSI.</li>
	<li>Place a dish of rowed embryos onto the microscope stage and have all embryos in field of view. Orientate the dish such that the scored line is on the side closest to the needle. Position the needle at an approximately 60&#176; angle so the tip is near the middle of the line and just above the surface of the water.</li>
	<li>Focus on the scored line and lower the needle until the tip comes into focus. Lower the tip a little more and break it open by running it gently into the ridges of the scored line. Raise the tip off the surface of the dish and position it at the top of the first row of zygotes and focus on the zygotes.</li>
	<li>Lower the tip such that it will impale the center of zygote coming in from around a 60&#176; angle. The needle easily penetrates P. miniata zygotes. Step on the foot pedal (or other means of forcing material from the injection needle) to introduce a bolus of injection solution into the embryo.</li>
	<li>Aim for a bolus that is 20-30% the diameter of the embryo, or between 25 and 80 pl. If the bolus is larger or smaller than this, adjust the duration of injection on the picospritzer and inject the next embryo. Continue to adjust the duration to obtain the desired bolus size. Start at 100 msec duration. Re-break the needle if greater than around 200 msec is needed to introduce the appropriate bolus. Discard the needle and prepare a new one if less than 15-20 msec is needed to obtain this bolus size as the needle size is too large and may disrupt normal development.</li>
	<li>Proceed to inject the remaining zygotes on the injection dish. Embryos can readily be injected until cleavage starts and into single blastomeres after first cleavage. Periodically lift the needle above the surface of the water to remove zygotes that stick onto the needle. If necessary re-break the tip of the needle or load a new needle if a blockage occurs.</li>
</ol>
5. Collecting Injected Embryos for Downstream Analysis
<ol>
	<li>Allow the embryos to develop at 15 &#176;C in SW. Maintain high surface area to volume in the cultures to allow for oxygen exchange. Ensure embryos are not crowded; keep cultures at or below 200 embryos/ml. Depending upon the desired stage of embryo, plan to collect injected embryos at around 20-24 hr&#160;post-fertilization. 
	NOTE: This is often an ideal time to sort and collect and because normally developing embryos are readily distinguished morphologically, but are not yet swimming.</li>
	<li>If embryos have hatched, gently salt shock to prevent swimming. Add 200 &#956;l of 5 M NaCl to 10 ml of SW in the injection dish while gently swirling the SW. After a few minutes, the embryos will fall to the bottom of the dish. Collect these and move them to normal salinity SW.</li>
	<li>Collect injected embryos under an appropriate fluorescent light source compatible with the tracer used in the injection solution. If no tracer was used, sort embryos to select for normal morphology.</li>
	<li>Using a mouth pipette, draw fluorescing embryos up with a minimal amount of SW and blow them out into a small culture dish filled with SW. Ideal mouth pipettes have a large enough opening for embryos to be pulled in and out without damaging them, but not so large that undesired embryos and extraneous SW are also pulled up.</li>
	<li>Once all of the desired embryos have been collected into the new dish, observe the embryos under fluorescent light and remove any un-injected embryos or poorly developed embryos.</li>
	<li>Culture sorted embryos to desired stages in an incubator and proceed with imaging or other desired protocols.</li>
</ol>

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No conflicts of interest declared by the authors.

There are two critical steps that are difficult for novice users of this technique but are essential for successfully creating morphant embryos. The first is selecting healthy oocytes that will mature and fertilize properly. The percentage of normal development in a culture depends on the season, the health of the animal, and the number of times that oocytes have been harvested from a single individual. Oocytes tend to be of better quality from April through October. It is important to look carefully at the oocytes befor...

The sea star, Patiria miniata, (commonly known as the bat star) is emerging as an interesting and versatile model system for a variety of cellular1-3, developmental4,5, evolutionary6-8, and ecological studies9-11. Adult P. miniata are distributed along the pacific coast from Sitka, Alaska to Baja, California12 and are readily maintained in marine aquaria. Oocytes are obtainable year round and each female can shed tens of thousands of eggs. Oocytes are easily matured and fertilized externally13. The resulting embryos are transparent allowing for easy observation; they develop synchronously, and require only sea water for development. Whole genome assembly and multiple transcriptomes are also available for P. miniata (<a href="http://www.echinobase.org/Echinobase/">Echinobase.org</a>). Such advantages make them ideal for a range of research and teaching purposes.
In recent years, P. miniata has become a model system for developmental gene regulatory network analyses14-16. The aim of such studies is to identify the entire compliment of regulatory genes and determine the network of their interactions. Much of this work entails perturbing gene expression through introduction of antisense oligonucleotides or in vitro synthesized mRNAs. Additionally, cis regulatory analyses are used to characterize the function of regulatory DNA15. These analyses require introduction of perturbation reagents and/or DNA reporter constructs into embryos. Furthermore, to characterize the downstream effects of these perturbations, one must assay many embryos for changes in gene expression of potential targets. Techniques for microinjection of many hundreds of zygotes are central for this work.
Echinoderms, including P. miniata, require many months to reach sexual maturity. Because of this, it is generally not practical to develop and maintain transgenic lines of these animals for experimentation. Therefore, breeding of transgenic adults cannot efficiently create modified embryos. Instead, perturbation must occur de novo through microinjection. Microinjection offers an opportunity to modify embryos with reagents that are not cell-permeable. The following protocol describes a method to introduce DNA, mRNA, cell tracers, and morpholino antisense oligonucleotides into hundreds of fertilized eggs in one 2-3 hr sitting through microinjection. This produces sufficient material for a variety of downstream experiments including, but not limited to, qPCR, in situ hybridization, RNA-Seq, and western blotting.

<table><tbody><tr><td>1-Methyladenine</td><td>Acros Organics (Fisher Scientific)</td><td>AC20131-1000</td><td></td></tr><tr><td>190 micron nitex nylon filter</td><td>Small Parts (originally Sefar)</td><td>CMN-0185-C/5PK-05</td><td></td></tr><tr><td>100 micron nitex nylon filter</td><td>Small Parts (originally Sefar)</td><td>CMN-0105-C/5PK-05</td><td></td></tr><tr><td>Polystyrene Petri dishes, 60 mm x 15 mm</td><td>Fisher Scientific</td><td>FB0875713A</td><td></td></tr><tr><td>Capillary tubing</td><td>FHC, Inc</td><td>30-30-0</td><td>For pulling microinjection needles</td></tr><tr><td>Model P-97 Needle Puller</td><td>Sutter Instruments</td><td>P-97</td><td></td></tr><tr><td>Dextran, Rhodamine Green</td><td>Life Technologies</td><td>D7163</td><td>If injecting a GFP expression reporter, it is helpful to substitute Texas Red dextran as an injection tracer</td></tr><tr><td>Instant Ocean Sea Salt</td><td>Doctors Foster and Smith</td><td>CD-116528</td><td>Also available in many pet stores</td></tr><tr><td>Microloader tips</td><td>Eppendorf</td><td>5242 956.003</td><td></td></tr></tbody></table>

a method for microinjection of patiria minata zygotes

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and evolution of developmental processes. The sea star, Patiria miniata, is gaining prevalence as a model system for these types of studies which were previously performed almost exclusively in the sea urchins, Strongylocentrotus purpuratus and Lytechinus variegatus. An advantage of these model systems is the ease of producing modified embryos in which a particular gene is up or downregulated, labeling a group of cells, or introducing a reporter gene. A single microinjection method is capable of creating a wide variety of such modified embryos. Here, we present a method for obtaining gametes from P. miniata, producing zygotes, and introducing perturbing reagents via microinjection. Healthy morphant embryos are subsequently isolated for quantitative and qualitative studies of gene function. The availability of genome and transcriptome data for this organism has increased the types of studies that are performed and the ease of executing them.

The goal of this protocol is to introduce reagents into embryos. We demonstrate the effectiveness of the protocol by injecting a DNA reporter construct that drives the expression of green fluorescent protein (GFP). Injected embryos express GFP in clonal patches (Figure 4A-B) as the DNA incorporates during early cleavage. Many reagents that are desirable to introduce into embryos are toxic in high quantities and to suboptimal batches of embryos. Toxicity manifests by delaying development, arresting develo...

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A Method for Microinjection of Patiria minata Zygotes

Methods that produce morphant embryos are essential to study developmental mechanisms and gene regulatory networks. The sea star Patiria miniata is an emerging model system for these studies. Here we present a protocol for obtaining gametes, producing cultures of embryos, and rapid microinjection of zygotes from this species.

A Method for Microinjection of Patiria minata Zygotes

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and ...

a-method-for-microinjection-of-patiria-minata-zygotes

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Biology

10.9K Views.  Carnegie Mellon University.  Methoden die morphant embryo's zijn essentieel voor ontwikkelingsstoornissen mechanismen en gen-regulatorische netwerken te bestuderen. De zee ster Patiria miniata is een opkomende modelsysteem voor deze studies. Hier presenteren we een protocol voor het verkrijgen van gameten, producerende culturen van embryo's, en snelle micro-injectie van zygoten van deze soort. 

Video: A Method for Microinjection of Patiria minata Zygotes

Laser-assisted Cytoplasmic Microinjection in Livestock Zygotes

Cytoplasmic microinjection into one-cell embryos is a very powerful technique. As an example, it enables the delivery of genome editing tools that can create genetic modifications that will be present in every cell of an adult organism. It can also be used to deliver siRNA, mRNAs or blocking antibodies to study gene function in preimplantation embryos. The conventional technique for microinjecting embryos used in rodents consists of a very thin micropipette that directly penetrates the plasma membrane when advanced into the embryo. When this technique is applied to livestock animals it usually results in low efficiency. This is mainly because in contrast to mice and rats, bovine, ovine, and porcine zygotes have a very dark cytoplasm and a highly elastic plasma membrane that makes visualization during injection and penetration of the plasma membrane hard to achieve. In this protocol, we describe a suitable microinjection method for the delivery of solutions into the cytoplasm of cattle zygotes that has proved to be successful for sheep and pig embryos as well. First, a laser is used to create a hole in the zona pellucida. Then a blunt-end glass micropipette is introduced through the hole and advanced until the tip of the needle reaches about 3/4 into the embryo. Then, the plasma membrane is broken by aspiration of cytoplasmic content inside the needle. Finally, the aspirated cytoplasmic content followed by the solution of interest is injected back into the embryonic cytoplasm. This protocol has been successfully used for the delivery of different solutions into bovine and ovine zygotes with 100% efficiency, minimal lysis, and normal blastocysts development rates.

This protocol shows how to perform cytoplasmic microinjection in farm animal zygotes. This technique can be used to deliver any solution into the one-cell embryo such as genome editing tools to generate knockout animals.

Cytoplasmic microinjection into one-cell embryos is a very powerful technique. As an example, it enables the delivery of genome editing tools that can ...

Genetics

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

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

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

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

Microinjection of Zebrafish Embryos to Analyze Gene Function

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

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

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

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 ...

Developmental Biology

Microinjection Method for Anopheles gambiae Embryos

Embryo microinjection techniques are essential for many molecular and genetic studies of insect species. They provide a means to introduce exogenous DNA fragments encoding genes of interest as well as favorable traits into the insect germline in a stable and heritable manner. The resulting transgenic strains can be studied for phenotypic changes resulting from the expression of the integrated DNA to answer basic questions or used in practical applications. Although the technology is straightforward, it requires of the investigator patience and practice to achieve a level of skill that maximizes efficiency. Shown here is a method for microinjection of embryos of the African malaria mosquito, Anopheles gambiae. The objective is to deliver by microinjection exogenous DNA to the embryo so that it can be taken up in the developing germline (pole) cells. Expression from the injected DNA of transposases, integrases, recombinases, or other nucleases (for example CRISPR-associated proteins, Cas) can trigger events that lead to its covalent insertion into chromosomes. Transgenic An. gambiae generated from these technologies have been used for basic studies of immune system components, genes involved in blood-feeding, and elements of the olfactory system. In addition, these techniques have been used to produce An. gambiae strains with traits that may help control the transmission of malaria parasites.

Microinjection Method for Anopheles gambiae Embryos

Microinjection techniques are essential to introduce exogenous genes into the genomes of mosquitoes. This protocol explains a method used by the James laboratory to microinject DNA constructs into Anopheles gambiae embryos to generate transformed mosquitoes.

Embryo microinjection techniques are essential for many molecular and genetic studies of insect species. They provide a means to introduce exogenous DNA ...

Microinjection of Medaka Embryos for use as a Model Genetic Organism

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1, as in zebrafish and the mouse. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions.
To take advantage of medaka fish whose embryos are transparent and develop externally, microinjection is an essential technique to inject cell-tracers for labeling cells, mRNAs or anti-sense oligonucleotides for over-expressing and knocking-down genes of interest, and DNAs for making transgenic lines.

Medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions. This protocol highlights the key points for successful microinjection into medaka embryos, an important technique for embryological and genetic analysis using medaka and zebrafish in a laboratory.

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows ...

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (&#62;1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

The present protocol describes zygote microinjection of CRISPR-Cas9 and donor DNA to efficiently produce gene cassette knock-in and floxed mice.

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily ...

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression analyses by RNA interference and stable transgenesis remain limited in many species due to the particular anatomy and molecular cellular biology of the organism. For example, outside of the crown group Caenorhabditis that includes Caenorhabditis elegans3, stably transmitted transgenic lines in non-Caenorhabditis species have not been reported in this specious phylum (Nematoda), with the exception of Strongyloides stercoralis4 and Pristionchus pacificus5. To facilitate the expanding role of P. pacificus in the study of development, evolution, and behavior6-7, we describe here the current methods to use microinjection for making transgenic animals and gene knock down by RNAi. Like the gonads of C. elegans and most other nematodes, the gonads of P. pacificus is syncitial and capable of incorporating DNA and RNA into the oocytes when delivered by direct microinjection. Unlike C. elegans however, stable transgene inheritance and somatic expression in P. pacificus requires the addition of self genomic DNA digested with endonucleases complementary to the ends of target transgenes and coinjection markers5. The addition of carrier genomic DNA is similar to the requirement for transgene expression in Strongyloides stercoralis4 and in the germ cells of C. elegans. However, it is not clear if the specific requirement for the animals' own genomic DNA is because P. pacificus soma is very efficient at silencing non-complex multi-copy genes or that extrachromosomal arrays in P. pacificus require genomic sequences for proper kinetochore assembly during mitosis. The ventral migration of the two-armed (didelphic) gonads in hermaphrodites further complicates the ability to inject both gonads in individual worms8. We also demonstrate the use of microinjection to knockdown a dominant mutant (roller,tu92) by injecting double-stranded RNA (dsRNA) into the gonads to obtain non-rolling F1 progeny. Unlike C. elegans, but like most other nematodes, P. pacificus PS312 is not receptive to systemic RNAi via feeding and soaking and therefore dsRNA must be administered by microinjection into the syncitial gonads. In this current study, we hope to describe the microinjection process needed to transform a Ppa-egl-4 promoter::GFP fusion reporter and knockdown a dominant roller prl-1 (tu92) mutant in a visually informative protocol.

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

In model organisms, transgenesis can manipulate gene functions while RNAi can knockdown specific mRNA transcripts 1-2. This protocol aims to illustrate the techniques needed to introduce stably transmitted DNA and transient double stranded RNA into the necromenic nematode Pristionchus pacificus for studies in evolutionary, developmental, and behavioral biology.

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression ...

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 ...

Zebrafish Microinjection Techniques

One of the major advantages to working with zebrafish (Danio rerio) is that their genetics can be easily manipulated by microinjection of early stage embryos. Using this technique, solutions containing genetic material or knockdown constructs are delivered into the blastomeres: the embryonic cells sitting atop the yolk of the newly fertilized egg. Delivery into the cytoplasm is achieved either through direct injection into the blastomere, or via natural cytoplasmic movements that occur after a solution is injected into the yolk. Successful genetic manipulations are usually followed by quantification of embryonic phenotypes in order to elucidate the genetic mechanisms of development.
This video will provide an introduction to carrying out microinjections in zebrafish embryos. The discussion begins with a review of the essential tools for the technique, including the injection apparatus and the microinjector, which controls fluid movement with pressure pulses of air. Next, important preparatory steps are demonstrated, such as the pouring of agar plates to stabilize embryos during injection and calibration of the microinjection apparatus. The injection procedure is then presented along with tips on when and where injections should be performed. Finally, applications of the microinjection technique are discussed, including gene overexpression via mRNA injection, gene silencing by delivery of antisense morpholino oligonucleotides, and the generation of transgenic zebrafish using specially engineered plasmid DNA.

Introduction to Microinjection of Early Zebrafish Embryos

One of the major advantages to working with zebrafish (Danio rerio) is that their genetics can be easily manipulated by microinjection of early stage ...

A Method for Microinjection of Patiria minata Zygotes

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