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 border="0" cellpadding="0" cellspacing="0" width="648">
	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Name of Material/ Equipment</td>
			<td style="width:147px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">1-Methyladenine</td>
			<td style="width:147px;">Acros Organics (Fisher Scientific)</td>
			<td style="width:119px;">AC20131-1000</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">190 micron nitex nylon filter</td>
			<td style="width:147px;">Small Parts (originally Sefar)</td>
			<td style="width:119px;">CMN-0185-C/5PK-05</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">100 micron nitex nylon filter</td>
			<td style="width:147px;">Small Parts (originally Sefar)</td>
			<td style="width:119px;">CMN-0105-C/5PK-05</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Polystyrene Petri Dishes, 60mm x 15mm</td>
			<td style="width:147px;">Fisher Scientific</td>
			<td style="width:119px;">FB0875713A</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Capillary tubing</td>
			<td style="width:147px;">FHC, Inc</td>
			<td style="width:119px;">30-30-0</td>
			<td style="width:167px;">For pulling microinjection needles</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">Model P-97 Needle Puller</td>
			<td style="width:147px;">Sutter Instruments</td>
			<td style="width:119px;">P-97</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="101">
			<td height="101" style="height:101px;width:216px;">Dextran, Rhodamine Green</td>
			<td style="width:147px;">Life Technologies</td>
			<td style="width:119px;">D7163</td>
			<td style="width:167px;">If injecting a GFP expression reporter, it is helpful to substitute Texas Red dextran as an injection tracer</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Instant Ocean Sea Salt</td>
			<td style="width:147px;">Doctors Foster and Smith</td>
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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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JoVE Journal

Biology

10.9K Views.  Carnegie Mellon University. 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.

Video: A Method for Microinjection of Patiria minata Zygotes

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 ...

Generation of Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

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 ...

Analysis of mRNA Nuclear Export Kinetics in Mammalian Cells by Microinjection

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although the nuclear export of mRNA has been studied extensively in Xenopus oocytes1 and genetically tractable organisms such as yeast2 and the Drosophila derived S2 cell line3, few studies had been conducted in mammalian cells. Furthermore the kinetics of mRNA export in mammalian somatic cells could only be inferred indirectly4,5. In order to measure the nuclear export kinetics of mRNA in mammalian tissue culture cells, we have developed an assay that employs the power of microinjection coupled with fluorescent in situ hybridization (FISH). These assays have been used to demonstrate that in mammalian cells, the majority of mRNAs are exported in a splicing dependent manner6,7, or in manner that requires specific RNA sequences such as the signal sequence coding region (SSCR) 6. In this assay, cells are microinjected with either in vitro synthesized mRNA or plasmid DNA containing the gene of interest. The microinjected cells are incubated for various time points then fixed and the sub-cellular localization of RNA is assessed using FISH. In contrast to transfection, where transcription occurs several hours after the addition of nucleic acids, microinjection of DNA or mRNA allows for rapid expression and allows for the generation of precise kinetic data.

Here we describe an assay that employs the power of microinjection coupled with fluorescent in situ hybridization in order to accurately measure the nuclear export kinetics of mRNA in mammalian somatic cells.

In eukaryotes, messenger RNA (mRNA) is transcribed in the nucleus and must be exported into the cytoplasm to access the translation machinery. Although ...

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

Single-cell Microinjection for Cell Communication Analysis

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a hemichannel by each neighboring cell to form the gap junction. In mammalian cells, the hemichannel is formed by six connexins, monomers with four transmembrane domains and a C and N terminal within the cytoplasm. Gap junctions permit the exchange of ions, second messengers, and small metabolites. In addition, they have important roles in many forms of cellular communication within physiological processes such as synaptic transmission, heart contraction, cell growth and differentiation. We detail how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells. It is expected that in functional gap junctions, the dye will diffuse from the loaded cell to the connected cells. It is a very useful technique to study gap junctions since you can evaluate the diffusion of the fluorescence in real time. We discuss how to prepare the cells and the micropipette, how to use a micromanipulator and inject a low molecular weight fluorescent dye in an epithelial cell line.

We describe here how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells, and provide some useful tips. We expect that this paper will help everyone to evaluate the degree of cellular coupling due to functional gap junctions. Everything described here could be, in principle, adapted to other fluorescent dyes with molecular weight below 1,000 Daltons.

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a ...

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 ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...