We thank Santiago Suarez for critical reading of the manuscript and Betty Cowgill for aid in photography. We also thank the anonymous reviewers for their critical feedback. This work is supported by the University of Delaware Research Fund.

1. Preparation of Protamine Sulfate (PS) Coated Dishes
<ol>
	<li>Prepare 1% solution of protamine sulfate (PS) by adding 0.5 g of PS to 50 ml of deionized, distilled water (ddH2O) in a 50 ml conical tube. Shake well at high speed on a bench shaker at room temperature for 1-2 hr to ensure complete dissolution of PS. This solution can be stored at 4 &#176;C for at least 3 months (make sure to completely dissolve gel-like precipitate before each use)3.</li>
	<li>Take a sleeve of 60 mm x 15 mm polystyrene Petri dishes and lay out both lids and bottoms on the bench.</li>
	<li>Pour 1% PS solution in each dish (both bottoms and lids can be used) just enough to cover the surface, leave for at least 2 min. The leftover PS solution can be reused many times within 3 months when stored at 4 &#176;C.</li>
	<li>Place PS-treated dishes in a beaker filled with distilled water (dH2O). Leave the beaker under running dH2O for at least 10 min.</li>
	<li>PS-coated dishes can be used immediately or air dry for storage. Cover them to prevent dust accumulation. They can be stored at room temperature for 1 month3.</li>
</ol>
2. Obtain Sea Urchin Gametes and Immobilize the Eggs on a PS-coated Dish for Microinjection
<ol>
	<li>Spawn sea urchins by shaking or intracoelomic injection of up to 1 ml of 0.5 % KCl to induce contraction of smooth muscles to release gametes3&#160;(Figure 2).</li>
	<li>If the animal is a male, as indicated by the white color sperm, collect sperm dry from the surface of an animal into a 1.5 ml tube. Dry sperm can be stored at 4 &#176;C for a week without significant loss of biological function.</li>
	<li>If the animal is a female, as indicated by the yellow color due to the egg yolk content, collect its gametes by immersing the gonopores in a beaker full of sea water. The eggs will be released from gonopores and settle down by gravity.</li>
	<li>Filter the eggs from debris such as animal spines using 80 &#956;m Nylon filter mesh. The best results are achieved if the eggs are used within 5-7 hr after spawning.</li>
	<li>Dejelly the eggs:
	<ol>
		<li>Prepare 100 ml of acidic sea water for approximately 1 ml of egg. Use 0.5 M citric acid to adjust pH of sea water from pH 8.0 to 5.1-5.2. Too low of a pH will decrease fertilization and too high of a pH will not allow eggs to attach to the PS coated plates.</li>
		<li>Add eggs (not more than 1 ml) to acidic sea water and allow the eggs to settle down on ice for 10 min. Alternatively, the removal of the jelly coat can be facilitated by gentle swirling of the eggs during the treatment. One can effectively dejelly S. purpuratus eggs in 3-4 min this way.</li>
		<li>Carefully pour off acidic sea water, add fresh sea water and let the eggs settle down on ice again. Dejellied eggs can be stored on ice for up to 6 hr without fertilization problems</li>
	</ol>
	</li>
</ol>
3. Row the Eggs
<ol>
	<li>Prepare 1 M stock solution of 3-aminotriazole (3-AT, MW = 84.08) by dissolving 0.84 g of 3-AT in 10 ml of ddH2O. This solution can be stored at 4 &#176;C for up to 6 months.</li>
	<li>Prepare 1 mM working solution of 3-AT by adding 50 &#956;l of 1 M stock solution to 50 ml of prechilled sea water. Keep the solution on ice.</li>
	<li>Prepare mouth pipette (Figure 3) for gentle handling of the eggs:
	<ol>
		<li>Pipette tip is manually pulled from glass micropipettes (100&#160;x OD 1.09 mm). Hold the micropipette with both hands over the open flame. As the glass begins to melt, carefully pull the ends of the pipette in opposing directions.</li>
		<li>Connect one pulled pipette to a plastic tubing (ID 1.14 mm), seal tight with Parafilm. The length of the tubing should be approximately 50-70 cm.</li>
		<li>Insert a sterile filtered P20 or P200 tip at the end of the tube opposite to the glass tubing to be used as the mouth piece.</li>
		<li>Clip the end of the glass micropipette with scissors to adjust the diameter of the tip. The optimal diameter slightly exceeds the diameter of the egg (80 &#956;m).</li>
	</ol>
	</li>
	<li>Pour &#160;&#160;4 ml of 1 mM 3-AT sea water into each PS-coated dish (bottom or lid).</li>
	<li>Take the mouth pipette, put the P20/P200 tip in the mouth and secure it with the teeth. To aspirate eggs, first aspirate a small amount of sea water. By having a column of liquid prior to loading the eggs, one will have maximal control over the mouth pipetting technique.</li>
	<li>Position the micropipette tip near the eggs and gently aspirate in several hundreds of eggs in the pipette. Confine the eggs within the glass micropipette.</li>
	<li>Row dejellied eggs in a straight line on a dish by gently blowing them out of the mouth pipette while moving the tip in a straight line. Row the eggs right before the injections, because fertilization problems may occur due to prolonged exposure to 3-AT sea water3. Moreover, cationic adhesives like PS can be toxic to eggs, and it is not advisable to expose embryos to these reagents for long periods, especially if the fertilization envelopes have been removed.</li>
	<li>Make a scratch on the dish near the line of rowed eggs using a glass pipette. This scratch will be important to break the injection needle to adjust the flow of solution as needed. Alternatively, the scratch can be made before pouring 3-AT sea water to the dish.</li>
</ol>
4. Microinjection of the Sea Urchin Zygotes
<ol>
	<li>Turn on microinjection station. Here we describe the use of FemtoJet injection system.</li>
	<li>Pull injection capillaries using a needle puller. An example of a good needle is depicted in Figure 4. Note: For RNA injections, injection capillaries should be pretreated to avoid RNAase contamination31:
	<ol>
		<li>Place injection capillaries into 50 ml conical tubes and add 50 ml of filtered methanol/HCl (9:1). Shake well and leave overnight for no longer than 10 hr.</li>
		<li>Pour off the methanol/HCl and rinse the capillaries 2x with filtered methanol in the same tube.</li>
		<li>Drain methanol as much as possible and lyophilize the capillaries overnight to ensure complete removal of methanol.</li>
	</ol>
	</li>
	<li>Carefully mount the needle capillary on a needle loading holder and load it with &#60;0.5 &#956;l of a sample solution using microloader tips. Sample solutions usually contain 20% glycerol, depending on the injected material3.</li>
	<li>Place the dish with rowed eggs on the microscope stage (eggs should be aligned vertically when viewed through eyepieces) and focus on the eggs.</li>
	<li>Carefully mount the loaded needle onto the needle microinjecting holder and adjust the position of the needle to have a perfectly focused tip in the middle of the field.</li>
	<li>Depress the button to apply the maximum pressure for needle cleaning. Solution should come out of the needle. Adjust the pressure if necessary: injection pressure should be in the range of 90-360 hPa, with the compensation pressure at approximately 1/3 of the injection pressure.</li>
	<li>Adjust the solution flow by injecting into unfertilized egg. Using a joystick micromanipulator, direct the tip of the injection needle to the unfertilized egg. To facilitate needle entry, create a slight trembling by gentle tapping of the stage with a hard object such as a screw driver. An injection bolus forming inside the egg should be seen (Figure 5).</li>
	<li>If the injection bolus is not visible, slightly raise the tip of the needle above the eggs, locate the scratch on the dish and adjust it to be positioned in the middle of the field. Gently tap the needle tip to the scratch mark to break the tip of the injection needle. Adjust the injection and compensation pressures to ensure that the injection bolus does not exceed 1/5 of the diameter of the egg (1-2 pm).</li>
	<li>Once the injection bolus is calibrated, fertilize the eggs by adding diluted sperm (1:1,000) or 1-2 &#956;l of sperm. Mix well carefully to prevent dislodging the row of eggs.</li>
	<li>Start injections immediately after the fertilization envelope becomes visible (Figure 5). Move along the line of rowed eggs using the stage controller and inject as many zygotes as possible by poking them with the needle controlled by joystick micromanipulator.</li>
	<li>After 10-15 min, the newly fertilized eggs will become hardened and cannot be injected any longer. Remove the dish from the stage and carefully aspirate out sperm in 3-AT sea water using a plastic Pasteur pipette.</li>
	<li>Do not let the zygotes dry. Quickly add fresh prechilled sea water to cover the dish. Incubate the embryos at 15 &#176;C.</li>
</ol>

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The authors declare&#160;no competing financial interests or other conflicts of interest.

Microinjection is a powerful technique for delivering various treatments such as DNA, mRNA, recombinant proteins, loss of function and gain of function reagents, dyes and their combinations into eggs, embryos, and cells of various organisms1-7. However, several considerations should be kept in mind when designing a microinjection experiment.
It is critically important to consider the solubility of the delivered treatment and the injection volume. If the microinjected solution tends ...

Efficient and reproducible treatment delivery is one of the main methodological challenges for researchers. Several methods have been established to transiently deliver treatment solutions into the eggs, embryos, and cells. These methods include electroporation (based on a generating transient pores in the membrane using short electrical pulses)1,2, lipofection (delivery through the fusion of treatment-containing liposomes with the membrane)1, microparticle bombardment1 (DNA is precipitated on the micron-sized metal particles that are then used to penetrate the cells at high velocity), and transduction (virus is used as a delivery vehicle of transgenes). At the moment, microinjection is the only approach that holds the advantage of delivering any solution with 100% efficiency with minimal reagents. Moreover, a single injection solution can be composed of a complex cocktail of treatments. This technique has been used to successfully microinject the eggs and embryos from numerous species such as sea urchins3,4, zebra fish5, mouse6, frog7, and cattle8 as well as single cells in the tissue culture9. Single blastomeres injections at later developmental stages have also been conducted10-12.
The current methods of microinjections are based on the pressure-injection method that was initially described by Hiramoto10; however, great progress has been made towards optimization of this process. Excellent microinjection techniques have been described elsewhere11, and here we describe one of the specific methods that is currently used to microinject sea urchin (Strongylocentrotus purpuratus) newly fertilized eggs. For over a century, sea urchins have been a valuable experimental model15,16. Sea urchins are evolutionarily closely related to chordates (including us) and analysis of their genome revealed that they contain all the major gene families as the human17. They produce a large number of synchronously developing transparent embryos that can be easily manipulated. Using sea urchin as a model organism, the sea urchin community has contributed to our understanding of the fertilization process18-21, cell biological processes22-24, and the gene regulatory networks (GRNs)25-28.
Microinjection into sea urchin zygotes requires several steps. First, the eggs need to be immobilized prior to injections (described below). Microinjection dishes are coated with protamine sulfate (PS), which creates a positively-charged surface to which the negatively charged eggs can adhere3. The eggs are dejellied by incubation in acidic sea water (pH 5.15) for 10 min, followed by two washes in natural sea water or artificial sea water (pH 8.0). The dejellied eggs are carefully rowed in a straight line in the middle of PS-coated dish in the presence of 1 mM 3-aminotriazole (3-AT), which is required to inhibit the activity of ovoperoxidase that is secreted from the cortical granules of the egg as a result of fertilization29. This step is important to prevent hardening of the fertilization envelope and to facilitate microinjection needle entry. As an alternative to 1 mM 3-AT, 10 mM paraminobenzoic acid (PABA) can be used. The injection solution is loaded into a microinjection needle using specialized microloading pipette tip and mounted on a holder attached to micromanipulator and pressure unit (Figure 1). Each needle can be used to microinject individual zygotes in multiple experiments on separate days. Microinjection can be performed for 10-15 min until the zygotes harden. The zygotes are then washed with the sea water and cultured at 15 &#176;C. When the embryos reach hatching blastula stage, they release hatching enzyme that digests components of the fertilization envelope30 and allow them to naturally detach from the PS-coated dish. If necessary, the embryos can be gently detached from the dish using a mouth pipette or Pasteur pipette by gently blowing sea water onto the embryos. The described method enables efficient and reliable microinjection of 100-400 newly fertilized eggs on a single dish, providing a high-throughput method for downstream analyses.

<table><tbody><tr><td>Glass Pasteur pipettes</td><td>VWR</td><td>14673-043</td><td>&amp;nbsp;</td></tr><tr><td>Inverted microscope&amp;nbsp; Axiovert 40 &amp;deg;C</td><td>Zeiss</td><td>4109431007990000</td><td>Injection microscope</td></tr><tr><td>Microloader tips</td><td>Eppendorf</td><td>5242 956.003</td><td>Load injection solution</td></tr><tr><td>Nylon filter mesh 80&amp;nbsp;&amp;mu;m</td><td>Amazon.com</td><td>03-80-37</td><td>Filter eggs to get rid of debris</td></tr><tr><td>P20 or P200 Aerosol Barrier Pipette Tips</td><td>Fisher Scientific</td><td>02707432 or 02707430</td><td>Part of a mouth pipette</td></tr><tr><td>Parafilm</td><td>Fisher Scientific</td><td>13 374 12</td><td>Part of a mouth pipette</td></tr><tr><td>Polyethylene tubing</td><td>Intramedic</td><td>PE-160</td><td>Part of a mouth pipette</td></tr><tr><td>Protamine sulfate</td><td>MP Biomedicals, LLC</td><td>194729</td><td>Attach dejellied eggs to injection dishes</td></tr><tr><td>Sea urchins &lt;em&gt;S. purpuratus&lt;/em&gt;</td><td>Pt. Loma Marine Invertebrate Lab</td><td>N/A</td><td>&amp;nbsp;</td></tr><tr><td>Sea water</td><td>any pet store</td><td>Instant Ocean</td><td>&amp;nbsp;</td></tr><tr><td>Sterile 60 mm x 15 mm Polystyrene Petri Dish</td><td>Fisher Scientific</td><td>0875713A</td><td>Injection dishes</td></tr><tr><td>Three-Axis Coarse Positioning Micromanipulator MMN-1</td><td>Narishige</td><td>9124</td><td>Manipulate injection needle</td></tr><tr><td>Three-Axis Joystick Type Oil Hydraulic Fine Micromanipulator MMO-202ND</td><td>Narishige</td><td>9212</td><td>Manipulate injection needle</td></tr><tr><td>Transfer pipettes</td><td>Fisher Scientific</td><td>13-711-9AM</td><td>&amp;nbsp;</td></tr><tr><td>Vertical&amp;nbsp; needle puller&amp;nbsp;</td><td>Narishige</td><td>PC-10</td><td>Pull injection needles</td></tr></tbody></table>

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.

GFP and mCherry reporter constructs were in vitro transcribed and microinjected into the newly fertilized eggs. Embryos were incubated at 15 &#176;C for 24 hr (until the blastula stage) and imaged using Zeiss Observer Z1 microscope. Injection of reporter constructs did not lead to any developmental defects (Figure 6). For quantification of fluorescent signals, image acquisition was performed at low magnification (100X) to maximally capture fluorescent pixels (Figures 6D-F). Fluo...

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High Throughput Microinjections of Sea Urchin Zygotes

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 ...

high-throughput-microinjections-of-sea-urchin-zygotes

Research

JoVE Journal

Biology

14.5K Views.  University of Delaware. 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.

Video: High Throughput Microinjections of Sea Urchin Zygotes

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

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

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 ...

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 ...

Mosaic Zebrafish Transgenesis for Evaluating Enhancer Sequences

The completion of the human genome sequence, along with that of many other species, has highlighted the challenge of ascribing specific function to non coding sequences. One prominent function carried out by the non coding fraction of the genome is to regulate gene transcription; however, there are no effective methods to broadly predict cis-regulatory elements from primary DNA sequence. We have developed an efficient protocol to functionally evaluate potential cis-regulatory elements through zebrafish transgenesis. Our approach offers significant advantages over cell-culture based techniques for developmentally important genes, since it provides information on spatial and temporal gene regulation. Conversely, it is faster and less expensive than similar experiments in transgenic mice, and we routinely apply it to sequences isolated from the human genome. Here we demonstrate our approach to selecting elements for testing based on sequence conservation and our protocol for cloning sequences and microinjecting them into zebrafish embryos.

We demonstrate our approach to finding potential enhancer elements from developmentally regulated genes and evaluating their function through mosaic zebrafish transgenesis.

The completion of the human genome sequence, along with that of many other species, has highlighted the challenge of ascribing specific function to non ...

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

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.

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.

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

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 ...

High Throughput Microinjections of Sea Urchin Zygotes

Summary

Explore More Videos

Chapters in this video

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

Laser-assisted Cytoplasmic Microinjection in Livestock Zygotes

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

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

Microinjection of Zebrafish Embryos to Analyze Gene Function

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Mosaic Zebrafish Transgenesis for Evaluating Enhancer Sequences

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

A Method for Microinjection of Patiria minata Zygotes

Zebrafish Microinjection Techniques